(PDF) In Vitro Embryogenesis in Higher Plants (1)

In Vitro

Embryogenesis

in Higher Plants

Maria Antonietta Germanà

Maurizio Lambardi Editors

Methods in

Molecular Biology 1359

M

ETHODS

IN

M

OLECULAR

B

IOLOGY

Series Editor

John M. Walker

School of Life and Medical Sciences

University of Hertfordshire

Hat ﬁ eld, Hertfordshire, AL10 9AB, UK

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In Vitro Embryogenesis

in Higher Plants

Edited by

Maria Antonietta Germanà

Dipartimento Scienze Agrarie e Forestali, Università degli Studi di Palermo, Palermo, Italy

Maurizio Lambardi

IVALSA/Trees and Timber Institute, National Research Council (CNR), Sesto Fiorentino, Florence, Italy

ISSN 1064-3745 ISSN 1940-6029 (electronic)

Methods in Molecular Biology

ISBN 978-1-4939-3060-9 ISBN 978-1-4939-3061-6 (eBook)

DOI 10.1007/978-1-4939-3061-6

Library of Congress Control Number: 2015952782

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Cover image: Somatic embr yogenesis in sweet orange (Citrus sinensis (L.) Osbeck). Photo of Maria Antonietta Germanà

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Editors

Maria Antonietta Germanà

Dipartimento Scienze Agrarie e Forestali

Università degli Studi di Palermo

Palermo , Italy

Maurizio Lambardi

IVALSA/Trees and Timber Institute

National Research Council (CNR)

Sesto Fiorentino, Florence , Italy

Dedication

To Emanuela,

Maria Luisa,

Antonio and

Gabriele,

the best experiments of my life

Maria Antonietta Germanà

To my beloved children

Matilde and Tommaso

Maurizio Lambardi

vii

I was fortunate to start my research career in plant tissue culture in the 1970s when this ﬁ eld

of research was expanding rapidly. The next few decades witnessed an exponential growth in

knowledge, understanding, and application of many tissue culture protocols to a wide range of

plant species. Then followed a period in the 1990s and turn of the century when plant tissue

culture research was neglected. Many of the leading researchers of the era such as Toshio

Murashige, Pierre Debergh, and Walter Preil retired. Postgraduate students and young

researchers now wanted to work in the new ﬁ eld of biotechnology. For a time, plant tissue

culture was becoming the “forgotten art” even though it underpinned new biotechnologies

such as plant transformation. However, some scientists continued to work on plant tissue cul-

ture and applied new molecular genetic techniques, such as gene identiﬁ cation, function, and

expression, to an understanding of basic plant pathways such as embryogenesis. It has been

encouraging for me, as I now reach retirement, to see the next generation of experienced plant

tissue culturists now ﬁ lling the ranks of the experts who have gone before. Maurizio Lambardi

and Maria Antonietta Germanà are two of those scientists who are renowned for their research

on plant tissue culture. I have known Maurizio both through his research and his contribution

to the International Society for Horticultural Science in his role as Chair of the Commission

Molecular Biology and In Vitro Culture. Maurizio is both an accomplished researcher and a

genuine person who is passionate about his ﬁ eld of research. Maria Antonietta Germanà is an

experienced researcher in gametic and somatic embryogenesis in fruit crops. I recommend

them as leaders in their ﬁ eld and ideal authors of this book on embryogenesis.

When I ﬁ rst started working on plant tissue culture in the early 1970s, very little was

known about embryogenesis. Why species had a predetermined genetic bias to regenerate

from callus by embryogenic or organogenic pathways was a mystery. Of the species that

were easy to tissue culture, why was carrot embryogenic and tobacco organogenic? In the

1980s, one of my Ph.D. supervisors advised me not to work on embryogenesis because it

appeared to depend on “phases of the moon.” The message was that experimental results

were inconsistent because of our lack of understanding; thus it was not recommended as a

topic for students who were facing a deadline and needed reliable and repeatable results.

However, our knowledge of embryogenesis has been greatly expanded in recent years. This

book represents a detailed overview of the current status of research on embryogenesis and

the advances that have been made by researchers who have worked on biotechnology and

in vitro culture. Thus the book contains chapters on “Recent advances on genetic and

physiological bases of in vitro somatic embryo formation,” “A central role of mitochondria

for stress-induced somatic embryogenesis;” “…What can we learn from proteomics?,”

“Genome-wide approaches and recent insights,” and “Microspore embryogenesis.” There

are chapters on somatic embryogenesis in a range of horticultural species, and an excellent

series of protocols for embryogenesis from a range of explants.

Foreword

viii

I would recommend this book to students, researchers, and those who have an interest

in plant tissue culture, and to those who may not realize the importance of knowledge of

this “forgotten art.”

President of the International Society Roderick Drew

for Horticultural Science (ISHS)

Leuven, Belgium

Foreword

ix

Embryogenesis in higher plants, one of the different routes of morphogenesis of the plant

kingdom, is a fascinating example of cellular totipotency. In fact, different kinds of plant

cells (somatic, gametic, nucellar, and fertilized egg cells) are able to regenerate, in nature or

in vitro, an entire organism through the formation of a somatic, gametic, or zygotic embryo,

a bipolar structure without vascular connection with the surrounding tissue. In vitro

somatic, gametic, and zygotic embryogenesis, apomixis, and secondary embryogenesis are

actually valuable tools to support plant breeding, propagation, and conservation, with rel-

evant implications to agriculture, forestry, horticulture, and preservation of plant genetic

resources. Advances in plant biotechnology, and particularly in tissue culture, led in time to

a better understanding of the physiological and biochemical bases regulating the process of

plant embryogenesis, and to the establishment of more and more efﬁ cient protocols of

in vitro embryo induction, maturation, and conversion to plant. Moreover, the recent

molecular, genomic, and proteomic studies have produced additional valuable contribu-

tions to the comprehension of the in vitro embryogenic developmental process.

The intent of the book is to present an overview of recent advances, innovative applica-

tions, and future prospects of in vitro embryogenesis in higher plants by means of topical

reviews and stepwise protocols of selected species. With this goal, the book has been divided

into ﬁ ve parts. Part I contains reviews on general topics (microspore, zygotic and somatic

embryogenesis, in vitro and in vivo asexual embryogenesis, advances on the genetic, physi-

ological, and proteomic knowledge of somatic embryo formation, role of programmed cell

death and mitochondria in somatic embryogenesis, and innovation in the use of bioreac-

tors). The remaining part of the book contains stepwise protocols on somatic embryogen-

esis in selected horticultural plants ( Part II ) and forest trees ( Part III ), on gametic

embryogenesis ( Part IV ), and on some pivotal topics ( Part V ), such as the detection of

epigenetic modiﬁ cations during microspore embryogenesis, the in vitro embryogenesis and

plant regeneration from isolated zygotes, the synthetic seed production, the induction and

maturation of somatic embryos, and the cryostorage of embryogenic cultures. Some useful

“Notes,” a peculiarity of the series “Methods in Molecular Biology,” complete all the step-

wise chapters, with additional information directly coming from the authors’ valuable daily

experience in the tissue culture laboratory.

We are extremely grateful to all the authors for providing such excellent contributions,

coming from their remarkable expertise on the different aspects of in vitro plant embryo-

genesis. It is our hope that this book will be a useful source of information and ideas for

plant tissue culturists, cell biologists, embryologists, horticulturists, and operators of com-

mercial nurseries. It is also our hope that it will attract students and young scientists toward

the fascinating world of in vitro embryogenesis in higher plants.

Palermo, Italy Maria Antonietta Germanà

Sesto Fiorentino, Florence, Italy Maurizio Lambardi

Pref ace

xi

Contents

Foreword. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vii

Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix

Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xv

PART I REVIEWS ON GENERAL TOPICS

1 A Comparison of In Vitro and In Vivo Asexual Embryogenesis . . . . . . . . . . . . 3

Melanie L. Hand , Sacco de Vries , and Anna M. G. Koltunow

2 Somatic Versus Zygotic Embryogenesis: Learning from Seeds . . . . . . . . . . . . . 25

Traud Winkelmann

3 Recent Advances on Genetic and Physiological Bases

of In Vitro Somatic Embryo Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47

Maria Maddalena Altamura , Federica Della Rovere ,

Laura Fattorini , Simone D’Angeli , and Giuseppina Falasca

4 Do Mitochondria Play a Central Role in Stress-Induced Somatic

Embryogenesis? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

Birgit Arnholdt-Schmitt , Carla Ragonezi , and Hélia Cardoso

5 Dying with Style: Death Decision in Plant Embryogenesis. . . . . . . . . . . . . . . . 101

Shuanglong Huang , Mohamed M. Mira

, and Claudio Stasolla

6 Somatic Embryogenesis in Broad-Leaf Woody Plants:

What We Can Learn from Proteomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117

Sandra I. Correia , Ana C. Alves , Paula Veríssimo , and Jorge M. Canhoto

7 Advances in Conifer Somatic Embryogenesis Since Year 2000 . . . . . . . . . . . . . 131

Krystyna Klimaszewska , Catherine Hargreaves , Marie-Anne Lelu-Walter ,

and Jean-François Trontin

8 Molecular Aspects of Conifer Zygotic and Somatic Embryo Development:

A Review of Genome-Wide Approaches and Recent Insights . . . . . . . . . . . . . . 167

Jean-François Trontin , Krystyna Klimaszewska , Alexandre Morel ,

Catherine Hargreaves , and Marie-Anne Lelu-Walter

9 Androgenesis in Solanaceae. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209

Jose M. Seguí-Simarro

10 Bioreactors for Plant Embryogenesis and Beyond . . . . . . . . . . . . . . . . . . . . . . 245

Liwen Fei and Pamela Weathers

PART II PROTOCOLS OF SOMATIC EMBRYOGENESIS

IN SELECTED IMPORTANT HORTICULTURAL PLANTS

11 Somatic Embryogenesis and Genetic Modification of Vitis. . . . . . . . . . . . . . . . 263

Sadanand A. Dhekney , Zhijian T. Li , Trudi N. L. Grant ,

and Dennis J. Gray

xii

12 Somatic Embryogenesis in Peach-Palm (Bactris gasipaes)

Using Different Explant Sources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279

Douglas A. Steinmacher , Angelo Schuabb Heringer , Víctor M. Jiménez ,

Marguerite G. G. Quoirin , and Miguel P. Guerra

13 Somatic Embryogenesis: Still a Relevant Technique

in Citrus Improvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

Ahmad A. Omar , Manjul Dutt , Frederick G. Gmitter ,

and Jude W. Grosser

14 Somatic Embryogenesis Induction and Plant Regeneration

in Strawberry Tree (Arbutus unedo L.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 329

João F. Martins , Sandra I. Correia , and Jorge M. Canhoto

15 Somatic Embryogenesis in Olive (Olea europaea L. subsp. europaea

var. sativa and var. sylvestris) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341

Eddo Rugini and Cristian Silvestri

16 Somatic Embryogenesis in Crocus sativus L. . . . . . . . . . . . . . . . . . . . . . . . . . . 351

Basar Sevindik and Yesim Yalcin Mendi

17 Somatic Embryogenesis in Lisianthus (Eustoma russellianum Griseb.) . . . . . . . 359

Barbara Ruffoni and Laura Bassolino

18 Somatic Embryogenesis in Two Orchid Genera

(Cymbidium, Dendrobium) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371

Jaime A. Teixeira da Silva and Budi Winarto

19 Somatic Embryogenesis of Lilium from Microbulb Transverse

Thin Cell Layers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387

Pablo Marinangeli

20 Somatic Embryogenesis and Plant Regeneration of Brachiaria brizantha. . . . . 395

Glaucia B. Cabral , Vera T. C. Carneiro , Diva M. A. Dusi ,

and Adriana P. Martinelli

PART III PROTOCOLS OF SOMATIC EMBRYOGENESIS

IN SELECTED FOREST TREES

21 Somatic Embryogenesis in Pinus spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405

Itziar Aurora Montalbán , Olatz García-Mendiguren ,

and Paloma Moncaleán

22 Somatic Embryogenesis of Abies cephalonica Loud. . . . . . . . . . . . . . . . . . . . . . 417

Jana Krajňáková and Hely Häggman

23 Somatic Embryogenesis in Horse Chestnut (Aesculus hippocastanum L.) . . . . . 431

Maurizio Capuana

24 Somatic Embryogenesis in Araucaria angustifolia (Bertol.)

Kuntze (Araucariaceae) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 439

Miguel P. Guerra , Neusa Steiner , Francine L. Farias-Soares ,

Leila do N. Vieira , Hugo P. F. Fraga , Gladys D. Rogge-Renner

,

and Sara B. Maldonado

Contents

xiii

PART IV PROTOCOLS OF GAMETIC EMBRYOGENESIS

IN SELECTED HIGHER PLANTS

25 Anther Culture in Eggplant (Solanum melongena L.). . . . . . . . . . . . . . . . . . . . 453

Giuseppe Leonardo Rotino

26 Anther Culture in Pepper (Capsicum annuum L.). . . . . . . . . . . . . . . . . . . . . . 467

Verónica Parra-Vega and Jose M. Seguí-Simarro

27 Microspore Embryogenesis Through Anther Culture

in Citrus clementina Hort. ex Tan. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475

Benedetta Chiancone and Maria Antonietta Germanà

PART V STEPWISE PROTOCOLS ON PIVOTAL TOPICS

28 Detection of Epigenetic Modifications During Microspore

Embryogenesis: Analysis of DNA Methylation Patterns Dynamics . . . . . . . . . . 491

Pilar S. Testillano and María Carmen Risueño

29 Embryogenesis and Plant Regeneration from Isolated Wheat Zygotes . . . . . . . 503

Jochen Kumlehn

30 From Somatic Embryo to Synthetic Seed in Citrus spp.

Through the Encapsulation Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 515

Maurizio Micheli and Alvaro Standardi

31 From Stress to Embryos: Some of the Problems for Induction

and Maturation of Somatic Embryos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 523

Sergio J. Ochatt and Maria Angeles Revilla

32 Cryotechniques for the Long-Term Conservation of Embryogenic

Cultures from Woody Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 537

Elif Aylin Ozudogru and Maurizio Lambardi

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 551

Contents

xv

Contributors

MARIA MADDALENA ALTAMURA • Department of Environmental Biology , Sapienza

University of Rome , Rome , Italy

ANA C. ALVES • Associação UC INPROPLANT – Investigação, Desenvolvimento

Tecnológico e Internacionalização , Paço das Escolas , Coimbra , Portugal

BIRGIT ARNHOLDT-SCHMITT • EU Marie Curie Chair ICAAM, IIFA , Universidade de

Évora , Évora , Portugal

LAURA BASSOLINO • Ornamental Plants Research Unit, Consiglio per la ricerca in agricoltura

e l’analisi dell’economia agraria (CREA), Sanremo, Imperia, Italy

GLAUCIA B. CABRAL • Embrapa Genetic Resources and Biotechnology , Parque Estação

Biológica , Brasília , DF , Brazil

JORGE M. CANHOTO • Center for Functional Ecology, Department of Life Sciences ,

University of Coimbra , Coimbra , Portugal

MAURIZIO CAPUANA • Istituto di Bioscienze e Biorisorse (IBBR) – CNR, sede Firenze , Sesto

Fiorentino , Firenze , Italy

HÉLIA CARDOSO • EU Marie Curie Chair , ICAAM, IIFA, Universidade de Évora , Évora ,

Portugal

VERA T. C. CARNEIRO • Embrapa Genetic Resources and Biotechnology , Parque Estação

Biológica , Brasília , DF , Brazil

BENEDETTA CHIANCONE • Dipartimento di Scienze degli Alimenti , Università degli Studi

di Parma , Parma , Italy

SANDRA I. CORREIA • Center for Functional Ecology, Department of Life Sciences, University of

Coimbra, Coimbra, Portugal

SIMONE D’ANGELI • Department of Environmental Biology , Sapienza University of Rome ,

Rome , Italy

FEDERICA DELLA ROVERE • Department of Environmental Biology , Sapienza University

of Rome , Rome , Italy

SADANAND A. DHEKNEY • Sheridan Research and Extension Center , University of Wyoming ,

Sheridan , WY , USA

DIVA M. A. DUSI • Embrapa Genetic Resources and Biotechnology , Parque Estação

Biológica , Brasília , DF , Brazil

MANJUL DUTT • Citrus Research and Education Center , University of Florida/IFAS ,

Lake Alfred , FL , USA

GIUSEPPINA FALASCA • Department of Environmental Biology , Sapienza University of Rome ,

Rome , Italy

FRANCINE L. FARIAS-SOARES • Graduate Program in Plant Genetic Resources,

Plant Developmental Physiology and Genetics Laboratory , Federal University of Santa

Catarina (UFSC) , Florianópolis , SC , Brazil

LAURA FATTORINI • Department of Environmental Biology , Sapienza University of Rome ,

Rome , Italy

LIWEN FEI • Department of Biology and Biotechnology , Worcester Polytechnic Institute ,

Worcester , MA , USA

xvi

HUGO P. F. FRAGA • Graduate Program in Plant Genetic Resources, Plant Developmental

Physiology and Genetics Laboratory , Federal University of Santa Catarina (UFSC) ,

Florianópolis , SC , Brazil

OLATZ GARCÍA-MENDIGUREN • NEIKER-TECNALIA, Centro de Arkaute, Vitoria-Gasteiz,

Spain

MARIA ANTONIETTA GERMANÀ • Dipartimento di Scienze Agrarie e Forestali , Università

degli Studi di Palermo , Palermo , Italy

FREDERICK G. GMITTER • Citrus Research and Education Center , University of Florida/

IFAS , Lake Alfred , FL , USA

TRUDI N. L. GRANT • Mid-Florida Research and Education Center , University of Florida/

IFAS , Apopka , FL , USA

DENNIS J. GRAY • Mid-Florida Research and Education Center , University of Florida/

IFAS , Apopka , FL , USA

JUDE W. GROSSER • Citrus Research and Education Center , University of Florida/IFAS ,

Lake Alfred , FL , USA

MIGUEL P. GUERRA • Graduate Program in Plant Genetic Resources, Plant Developmental

Physiology and Genetics Laboratory, Federal University of Santa Catarina (UFSC),

Florianópolis, SC, Brazil

HELY HÄGGMAN • Genetics and Physiology Department, University of Oulu, Oulu, Finland

MELANIE L. HAND • Commonwealth Scientiﬁ c and Industrial Research Organization

(CSIRO), Agriculture. Waite Campus, Urrbrae, South Australia

CATHERINE HARGREAVES • Scion , Rotorua , New Zealand

ANGELO SCHUABB HERINGER • UENF-Universidade Estadual do Norte Fluminense , Darcy

Ribeiro , RJ , Brazil

SHUANGLONG HUANG • Department of Plant Science , University of Manitoba , Winnipeg ,

Canada

VÍCTOR M. JIMÉNEZ • CIGRAS , Universidad de Costa Rica , San Pedro , Costa Rica

KRYSTYNA KLIMASZEWSKA • Laurentian Forestry Centre , Canadian Forest Service, Natural

Resources Canada , Québec , QC , Canada

ANNA M. G. KOLTUNOW • Commonwealth Scientiﬁ c and Industrial Research Organization

(CSIRO), Agriculture. Waite Campus, Urrbrae, South Australia

JANA KRAJŇÁKOVÁ • Department of Agriculture and Environmental Science , University

of Udine , Udine , Italy ; Genetics and Physiology Department, University of Oulu, Oulu,

Finland

JOCHEN KUMLEHN • Leibniz Institute of Plant Genetics and Crop Plant Research (IPK)

Gatersleben , Stadt Seeland/OT Gatersleben , Germany

MAURIZIO LAMBARDI • IVALSA/Trees and Timber Institute, National Research Council

(CNR), Sesto Fiorentino, Florence, Italy

MARIE-ANNE LELU-WALTER • INRA, UR 0588 Unité Amélioration , Génétique et

Physiologie Forestières , Ardon, Orléans Cedex 2 , France

ZHIJIAN T. LI • Mid-Florida Research and Education Center , University of Florida/IFAS ,

Apopka , FL , USA

SARA B. MALDONADO • Department of Biodiversity and Experimental Biology , University

of Buenos Aires , Buenos Aires , Argentina

PABLO MARINANGELI • Agronomy Department, Universidad Nacional del Sur. Centro de

Recursos Naturales Renovables de la Zona Semiárida (CERZOS), National Research Council

(CONICET), Bahia Blanca, Buenos Aires, Argentina

Contributors

xvii

ADRIANA P. MARTINELLI • University of São Paulo, CENA , Piracicaba , SP , Brazil

JOÃO F. MARTINS • Center for Functional Ecology, Department of Life Sciences , University

of Coimbra , Coimbra , Portugal

YESIM YALCIN MENDI • Department of Horticulture, Faculty of Agriculture, University of

Çukurova, Adana, Turkey

MAURIZIO MICHELI • Georgoﬁ li Academy of Florence , Florence , Italy

MOHAMED M. MIRA • Department of Botany, Faculty of Science , Tanta University , Tanta ,

Egypt

PALOMA MONCALEÁN • NEIKER-TECNALIA, Centro de Arkaute, Vitoria-Gasteiz, Spain

ITZIAR AURORA MONTALBÁN • NEIKER-TECNALIA, Centro de Arkaute, Vitoria-Gasteiz,

Spain

ALEXANDRE MOREL • INRA, UR 0588 Unité Amélioration , Génétique et Physiologie

Forestières , Ardon, Orléans Cedex 2 , France

SERGIO J. OCHATT • INRA, CR de Dijon , UMR 1347 Agroécologie , Dijon Cedex , France

AHMAD A. OMAR • Citrus Research and Education Center , University of Florida/IFAS ,

Lake Alfred , FL , USA ; Biochemistry Department, College of Agriculture , Zagazig

University , Zagazig , Egypt

ELIF AYLIN OZUDOGRU • IVALSA/Trees and Timber Institute, National Research Council

(CNR), Sesto Fiorentino, Florence, Italy

VERÓNICA PARRA-VEGA • COMAV - Universitat Politècnica de València. CPI,

Valencia , Spain

MARGUERITE G. G. QUOIRIN • Programa de Pós-Graduação em Agronomia , UFPR ,

Curitiba , PR , Brazil

CARLA RAGONEZI • EU Marie Curie Chair , ICAAM, IIFA, Universidade de Évora , Évora ,

Portugal

MARIA ANGELES REVILLA • Departamento Biología de Organismos y Sistemas, Instituto de

Biotecnología de Asturias, Universidad de Oviedo, Oviedo, Spain

MARÍA CARMEN RISUEÑO • Pollen Biotechnology of Crop Plants, Biological Research Centre ,

CIB-CSIC , Madrid , Spain

GLADYS D. ROGGE-RENNER • Department of Cell Biology, Embryology and Genetics,

Plant Cell Biology Laboratory , UFSC , Florianópolis , SC , Brazil ; Department of Biological

Sciences , University of Joinville Region , Joinville , SC , Brazil

GIUSEPPE LEONARDO ROTINO • Vegetable Crops Research Unit, Consiglio per la ricerca in

agricoltura e l’analisi dell’economia agraria (CREA), Montanaso Lombardo, Lodi, Italy

BARBARA RUFFONI • Ornamental Plants Research Unit, Consiglio per la ricerca e l’analisi

dell’economia agraria (CREA), Sanremo, Imperia, Italy

EDDO RUGINI • Department of Agriculture, Forests, Nature and Energy (DAFNE) ,

University of Tuscia , Viterbo , Italy

JOSE M. SEGUÍ-SIMARRO • COMAV - Universitat Politècnica de València. CPI, Camino de

Vera , Valencia , Spain

BASAR SEVINDIK • Department of Horticulture, Faculty of Agriculture, University of Çukurova,

Adana, Turkey

CRISTIAN SILVESTRI • Department of Agriculture, Forests, Nature and Energy (DAFNE) ,

University of Tuscia , Viterbo , Italy

ALVARO STANDARDI • Georgoﬁ li Academy of Florence , Florence , Italy

CLAUDIO STASOLLA • Department of Plant Science , University of Manitoba , Winnipeg ,

Canada

Contributors

xviii

NEUSA STEINER • Department of Botany , UFSC , Florianópolis , SC , Brazil

DOUGLAS A. STEINMACHER • Programa de Pós-Graduação em Agronomia , UFPR ,

Curitiba , PR , Brazil ; Vivetech Agrociências , Marechal Candido Rondon , PR , Brazil

JAIME A. TEIXEIRA DA SILVA • P.O. Box 7, Miki-cho Post Ofﬁ ce , Kagawa-ken , Japan

PILAR S. TESTILLANO • Pollen Biotechnology of Crop Plants, Biological Research Centre ,

CIB-CSIC , Madrid , Spain

JEAN-FRANÇOIS TRONTIN • FCBA, Pôle Biotechnologie et Sylviculture Avancée , Équipe

Génétique et Biotechnologie, Campus Forêt-Bois de Pierroton , Cestas , France

PAULA VERÍSSIMO • Centre for Neuroscience and Cell Biology, Department of Life Sciences ,

University of Coimbra , Coimbra , Portugal

LEILA DO N. VIEIRA • Graduate Program in Plant Genetic Resources, Plant Developmental

Physiology and Genetics Laboratory , Federal University of Santa Catarina (UFSC) ,

Florianópolis , SC , Brazil

SACCO DE VRIES • Department of Biochemistry , University of Wageningen , Wageningen ,

The Netherlands

PAMELA WEATHERS • Department of Biology and Biotechnology , Worcester Polytechnic

Institute , Worcester , MA , USA

BUDI WINARTO • Indonesian Ornamental Crops Research Institute (IOCRI) ,

Pacet- Cianjur , West Java , Indonesia

TRAUD WINKELMANN • Leibniz Universität Hannover , Institute of Horticultural

Production Systems , Hannover , Germany

Contributors

Part I

Reviews on General Topics

3

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_1, © Springer Science+Business Media New York 2016

Chapter 1

A Comparison of In Vitro and In Vivo Asexual

Embryogenesis

Melanie L. Hand , Sacco de Vries , and Anna M. G. Koltunow

Abstract

In plants, embryogenesis generally occurs through the sexual process of double fertilization, which involves

a haploid sperm cell fusing with a haploid egg cell to ultimately give rise to a diploid embryo. Embr yogenesis

can also occur asexually in the absence of fertilization, both in vitro and in vivo . Somatic or gametic cells

are able to differentiate into embryos in vitro following the application of plant growth regulators or stress

treatments. Asexual embryogenesis also occurs naturally in some plant species in vivo, from either ovule

cells as part of a process deﬁ ned as apomixis, or from somatic leaf tissue in other species. In both in vitro

and in vivo asexual embryogenesis, the embryo precursor cells must attain an embryogenic fate without

the act of fertilization. This review compares the processes of in vitro and in vivo asexual embryogenesis

including what is known regarding the genetic and epigenetic regulation of each process, and considers

how the precursor cells are able to change fate and adopt an embryogenic pathway.

Key words Adventitious embryony , Apomixis , Cell fate , Gametic embryogenesis , Kalanchoë ,

Parthenogenesis , Somatic embryo genesis

1 Introduction

Embryogenesis describes the development of a single cell into an

embryo. In plant embryogenesis there is no cell migration, so

embryo pattern formation and cell type speciﬁ cation is interrelated

with oriented cell division and expansion. Within sexual angio-

sperm plant species, embryogenesis usually occurs in vivo within

ﬂ oral organs during the events of seed formation. Formation of an

embryo can also occur via asexual pathways in seeds, from somatic

plant cells in vivo or be induced experimentally from somatic plant

explants or gametes in vitro.

This review describes and compares the processes of in vivo

and in vitro asexual embryogenesis including what is currently

understood regarding the molecular mechanisms underlying each

process.

4

2 Types of Embryogenesis

The most prevalent form of embryogenesis in plants occurs follow-

ing double fertilization in the female gametophyte (embryo sac)

found in the ovule of the ﬂ ower, which gives rise to the embryo and

endosperm compartments of the seed (Table

1 ; Fig. 1a ). Haploid

male and female gametes form in the anther and ovule, respectively,

via meiosis and subsequent mitosis [

1 , 2 ]. Double fertilization initi-

ates when the male pollen tube containing two sperm cells enters the

ovule. One haploid sperm cell fuses with the meiotically derived hap-

loid egg cell in the female gametophyte to form the single-celled

diploid zygote, which then undergoes cell division and pattern form-

ing events to give rise to the diploid embryo [

3 ]. The other haploid

sperm cell fuses with the diploid central cell nucleus of the embryo

sac, which initiates divisions to form triploid endosperm that pro-

vides resources to the developing embryo [

4 ]. Ovule tissues that

surround the embryo and endosperm contribute to the seed coat.

Evolutionary speaking, embryogenesis is a much older process

than seed formation and initially resulted from the fusion of two

homospores into the zygote, gradually evolving in present day het-

erospory [

5 , 6 ]. The zygote formed following fusion of parental

gametes is the ﬁ rst cell evident during sexual reproduction with a

competence for embryogenesis. In plants, an “embryogenic” state

is not only restricted to the zygote and in the following sections,

ways of attaining an embryogenic state other than via fertilization

will be discussed (Table

1 ).

Apomixis is a term describing a suite of developmental processes

resulting in the formation of an asexual seed . Characteristic fea-

tures of all apomicts include fertilization - independent formation of

an egg cell or another somatic ovule cell into an embryo, and the

development of functional endosperm in apomicts occurs either

with or without fertilization [

7 , 8 ]. As a result, plants germinating

2.1 Zygotic (Sexual)

Embryogenesis

2.2 Asexual

Embryogenesis

in Seeds: Apomixis

and Parthenogenesis

in Cereals

Table 1

Characteristics of each type of embryogenesis considered in this review

Type of embryogenesis Precursor cell

Mode of

embryogenesis

Ploidy of

embryo

Biological

environment

Zygotic Egg Sexual Diploid In vivo

Parthenogenesis Egg Asexual Diploid In vivo

Adventitious embryony Nucellar/integument Asexual Diploid In vivo

Somatic embryog enesis Somatic cells Asexual Diploid In vitro/in vivo

Gametic embryogenesis Egg/sperm Asexual Haploid In vitro

Melanie L. Hand et al.

5

from seeds derived via apomixis are genetically identical to the

maternal parent.

Apomixis has evolved independently ac ross different angio-

sperm plant families and genera many times, and has been docu-

mented in more than 120 angiosperm genera that belong to

approximately 40 families [

9 ]. Apomixis is genetically controlled

by dominant loci in studied species and is not prevalent in agro-

nomically important plants [

10 ]. Apomixis mechanisms are gener-

ally divided into two categories: gametophytic or sporophytic,

based upon the location of the precursor cell which develops into

the embryo. In gametophytic apomixis , the embryo develops with-

out fertilization (termed parthenogenesis ) from an egg cell found

inside an embryo sac that has formed mitotically without prior

meiosis, and is thus chromosomally unreduced (Table

1 ; Fig. 1b ).

Fig. 1 Asexual embryogenesis occurs in vivo and in vitro from different cell types. ( a ) Floral organs and leaves

are some of the source plant tissue for inducing embryogenesis in vitro. Asexual embryos also form in ovules

in vivo; ( b ) Parthenogenesis involves the development of a chromosomally reduced or unreduced egg cell ( yel-

low ) into an embryo without fertilization ; ( c ) Nucellar or integument cells ( red ) adjacent to an embryo sac

within the ovule develop into embryos through adventitious embryony ; ( d ) In vivo somatic embryo genesis is

known to occur in species such as Kalanchoë , where the embryos develop along leaf margins; ( e ) Gametic

embryogenesis involves the experimental induction of embryogenesis from gametic cells such as microspores

and ovules; ( f ) Embryogenesis can be induced in somatic cells following experimental treatment; ( g ) Embryos

formed via asexual embryogenesis may or may not possess a suspensor . At a heart-shaped stage, the typical

plant embryo contains precursor cells for the shoot apical meristem ( blue cells ), and the root apical meristem

which consists of a quiescent center ( orange cells ) and columella stem cells ( purple cells )

In Vitro and In Vivo Asexual Embryogenesis

6

Two common mechanisms termed diplospory and apospory give

rise to such embryo sacs. They are distinguished by whether the

starting cell is a megaspore mother cell or another somatic cell in

the ovule, respectively (see Hand and Koltunow [

7 ] for further

information). Gametophytic apomixis and parthenogenesis are

found and studied in species including eudicots Taraxacum ofﬁ ci-

nale (dandelion), Boechera spp., and Hieracium spp. and also in

grasses Pennisetum squamulatum and Paspalum simplex among

others [

11 – 14 ].

During sporophytic apomixis , which is also called adventitious

or nucellar embryony, embryos develop without fertilization

directly from diploid somatic ovule cells surrounding an embryo

sac (Table

1 ; Fig. 1c ). Most commonly, the embryos arise from

two different ovule tissues: the nucellus and the inner integument.

Nucellar embryony is widespread among Citrus species [

15 , 16 ].

The embryo initial cells that give rise to the asexual embryos dif-

ferentiate near the developing embryo sac [

17 ] and they can be

speciﬁ ed as early as the 2–4 nuclear stage of embryo sac formation

[

18 , 19 ]. The embryo initial cells develop and form globular-

shaped embryos that can only develop to maturity if the sexually

derived embryo sac is fertilized, as the sexual and asexual embryos

share the nutritive endosperm . The developing seed therefore con-

sists of one sexual embryo and one or more asexual embryos and is

termed polyembryonic. The sexually derived embryo may not

develop or survive germination [

17 ].

Asexual embryogenesis is evident within seeds of the “Salmon”

system of wheat . In contrast to gametophytic apomixis , a chromo-

somally reduced embryo sac develops via the usual events of meio-

sis, spore selection, and mitosis evident in sexually reproducing

angiosperms . However, salmon wheat lines are capable of up to

90 % parthenogenesis , whereby the egg is able to initiate embryo-

genesis without fertilization [

20 , 21 ]. Parthenogenesis capability

results from translocation of the short arm of wheat chromosome

1B with the short arm of chromosome 1R of rye. This particular

translocation results in the loss of two critical loci in wheat:

Suppressor of parthenogenesis ( Spg ) and Restorer of fertility ( Rfv1 ),

along with the gain of a Parthenogenesis ( Ptg ) locus from rye. In

addition to this translocation, parthenogenesis is dependent upon

organellar DNA from Aegilops causdata or A. kotschyi , demon-

strating the importance of cytoplasmic as well as nuclear factors in

asexual embryogenesis in vivo [

21 ]. The existence of fertilization-

independent embryo development from different cell types in the

ovules of apomicts suggests that multiple cells can acquire an

embryogenic state. This contrasts with sexual reproduction where

the embryogenic state is suppressed until fertilization and

restricted to the egg cell within the female gametophyte. In par-

thenogenetic cereals the embryogenic state is attained by the egg

Melanie L. Hand et al.

7

in the absence of fertilization whilst embryogenic competency is

suppressed in the remaining ovule cell types.

Somatic embryo genesis is known to occur in vivo in nature, where

embryos develop on the surface of plant tissue (Fig.

1d ) [ 22 ]. For

example, plants of the genus Kalanchoë reproduce asexually

through the ectopic formation of plantlets along their leaf margins

[

23 ]. The plantlets arise following proliferation of cells described

as “dormant meristems” that are found in notches along the leaf

margin [

24 , 25 ]. Some Kalanchoë species require stress to induce

plantlet formation while others do not and constitutively form

asexual plantlets. Because of this form of multiplication, Kalanchoë

species are known as “mother of thousands.” The embryo result-

ing from somatic embryo genesis is diploid and genetically identical

to the somatic precursor cells from which it was formed.

It is possible to induce asexual embryogenesis in vitro from gametic

cells including male microspores (termed androgenesis ), and from

egg cell s or the associated accessory cells found in the female game-

tophytes (termed gynogenesis) (Table

1 ; Fig. 1e ). This process

requires gametophytic cells to switch to a sporophytic embryo for-

mation pathway. Application of various stress treatments such as

cold/heat shock and starvation are applied to the anther, isolated

microspores, cultured ovules, ovaries, or ﬂ ower buds to induce the

switch [

26 – 28 ]. The resulting embryos are haploid , possessing

either maternal or paternal chromosomes depending on the game-

tophytic precursor cell. The production of haploid plants through

in vitro gametic embryogenesis is a powerful mechanism to gener-

ate homozygous lines much faster than using conventional breed-

ing. Colchicine induced chromosome doubling of haploid embryos

during, or just after, embryogenesis results in homozygous

doubled- haploid plants which are useful tools in trait discovery and

plant breeding applications [

29 ]. Currently, microspore embryo-

genesis is favored over gynogenesis as a mode of gametic embryo-

genesis because of its higher efﬁ ciency [

30 ].

In vitro somatic embryo genesis can also be induced in vegeta-

tive explants or cells following treatment with plant growth regula-

tor s ( PGR ) or stresses such as osmotic shock, dehydration, water

stress , and alteration of pH (reviewed in [

31 ]) (Fig. 1f ). A few stud-

ies have addressed correspondences and differences between zygotic

and somatic embryogenesis and suggest that the patterning and

speciﬁ cation events are quite similar [

32 ], with the exception of a

lack of the suspensor and dormancy in in vitro cultured somatic

embryos [

33 ]. Therefore, the most important step in vegetative

cells that undergo somatic embryogenesis must be to ﬁ rst gain the

“embryogenic” state. Recent work suggests that a release in sup-

pression of the embryogenic state is a plausible mechanism [

6 , 34 ].

2.3 Somatic

Embryogenesis In Vivo

from Leaves

2.4 In Vitro Somatic

and Gametic

Embryogenesis

In Vitro and In Vivo Asexual Embryogenesis

8

3 Attaining an Embryogenic State

A prerequisite for embryogenesis in plants is that the precursor cell

must attain an embryogenic state which provides the cellular com-

petence for embryo formation. During gametic embryogenesis ,

and gametophytic apomixis , the developing gametophyte cells

respond to induction signals that switch their fate from gameto-

phytic to sporophytic. During zygotic embryo genesis , the zygote

has acquired embryogenic competency following fertilization of

the egg cell . In somatic embryo genesis in vitro, and adventitious

embryony , the embryo precursor cells are somatic sporophytic cells

which ﬁ rst must attain the embryogenic state. Changing the devel-

opmental fate of a cell is therefore an important component of

both in vitro and in vivo asexual embryogenesis.

It has been proposed that somatic embryo genesis consists of

two distinct phases which are independent of each other and are

controlled by different factors [

35 ]. The initial stage is induction,

which involves the somatic cells attaining the embryogenic state

usually by the exogenous application of PGR. The following stage

is expression, where the newly differentiated embryonic cells

develop into an embryo without any further exogenous signals. It

is not yet known whether in vivo embryogenesis via adventitious

embryony similarly consists of two separate independent phases.

However, such a scenario could be envisaged where the sporo-

phytic ovule cells also ﬁ rst acquire embryonic competence by a

particular molecular signal, and then develop into an embryo with-

out fertilization via a separate developmental program.

In the process of in vitro somatic embryo genesis, somatic cells

attain the embryogenic state following the application of PGR .

Auxin is most commonly used [

36 ], although other PGR, includ-

ing cytokinin and abscisic acid, have proven capable of inducing

embryogenesis [

37 , 38 ]. Following treatment with PGR, the cells

are cultured on a hormone-free medium. Auxin plays major roles

in plant growth and morphogenesis including embryo sac develop-

ment and embryo patterning [

39 , 40 ]. In addition to treatment

with auxin, the frequency of somatic embryogenesis induction also

depends on the species, genotype, tissue, stage of development,

and endogenous hormone levels [

35 , 41 ]. Therefore although

auxin is a universal induction molecule, other factors must be

involved in the induction of embryonic competence. The role of

cellular stress response s in the induction of somatic embryogenesis

is increasingly being recognized. The process of culturing explants

for somatic embryogenesis involves wounding, sterilization, and

culturing of the explant, all which undoubtedly apply stress to the

cells involved. Furthermore, exogenous stresses such as osmotic,

heavy metal ion, temperature, and dehydration stresses can enhance

Melanie L. Hand et al.

9

somatic embryogenesis [ 42 – 46 ]. The induction of somatic

embryogenesis through the application of auxin or stresses may

imply an interaction between auxin and stress signaling . Auxin may

therefore activate a stress signaling response, which is involved in

inducing embryogenic competence . Many stress-related genes are

up-regulated during the early phases of somatic embryogenesis,

which supports this theory [

47 , 48 ].

Whether somatic cells in vitro and nucellar, integument cells

and unreduced egg cell s in apomicts in vivo acquire an embryo-

genic state via the same mechanism is currently unknown. Unlike

somatic embryo genesis, embryos formed through parthenogenesis

and adventitious embryony in apomicts are subject to the develop-

mental inﬂ uences of the ovule which may produce alternate cues

that induce an embryogenic state. Stress and alterations in ovule

pattern formation lead to a deregulation of apomixis in Hieracium

where embryos form ectopically in different ovule positions [

49 ].

Although no genes have yet been identiﬁ ed that are responsible for

inducing adventitious embryony, genes related to stress signaling

have been implied in the process of nucellar embryony in Citrus .

Kumar et al. [

50 ] used suppression subtractive hybridization (SSH)

and microarray to detect genes that were differentially expressed

during asexual embryo initiation and discovered genes related to

stress signaling, including heat shock proteins.

Some similarities exist in the morphology of the embryo pre-

cursor cell for in vitro somatic embryo genesis and in vivo adventi-

tious embryony . In Citrus species that undergo adventitious

embryony, those nucellar cells that ultimately differentiate into

embryos are distinguished from surrounding nucellar cells by their

large nuclei and dense cytoplasm [

51 ]. These nucellar initial cells

also have very thick callosic cell wall s and later become thinner

walled, rounder, larger, and with a prominent nucleus prior to cell

division [

17 ]. Histological observations of embryonic somatic cells

cultured in vitro from various species show that these embryonic

cells are relatively small and also contain large nuclei and dense

cytoplasm when compared to other somatic cells (reviewed in

Namasivayam [

52 ]). Large nuclei and dense cytoplasm are also

characteristic of cells that are precursors of the female gametophyte,

including the aposporous initial cell in aposporous apomictic plants,

distinguishing them from surrounding somatic cells [

1 , 53 ].

4 Embryo Morphology

Zygotic embryo genesis within angiosperms passes through a series

of sequential stages to give rise to the mature differentiated struc-

ture. In Arabidopsis and some other angiosperms, the ﬁ rst division

In Vitro and In Vivo Asexual Embryogenesis

10

of the zygote produces an apical cell that continues to be embryo-

genic, while the second basal cell is no longer embryogenic and

continues to form the multicelled suspensor . Further divisions of

the apical cell produce a globular embryo, and differentiation and

expansion of the cotyledons leads to heart and torpedo-shaped

embryos [

54 ]. Only the suspensor derived hypophyseal suspensor

cell continues to form the quiescent center and the columella stem

cells of the root meristem (Fig.

1g ) [ 55 ]. Variation in early cell

division patterning exists between different dicotyledonous spe-

cies, although the typical globular, heart, and torpedo morpho-

logical stages still usually occur [

54 ]. Zygotic embryogenesis in

monocotyledonous species differs from dicots mostly with respect

to planes of symmetry and the position of the shoot apical meri-

stem [

56 ]. Variation in embryo formation also exists between

monocot species. The embryo is the only plant structure in which

both the root and shoot apical meristem is formed simultaneously.

This requires a highly complex series of pattern forming and speci-

ﬁ cation events, including establishment of small populations of

stem cells. These cells continue to support the formation and activ-

ity of meristems during the remainder of the plant life cycle (for a

recent review see [

57 ]). Extensive studies have revealed molecular

details of the formation of the major tissue types as well as the

meristems themselves during embryogenesis [

6 ].

The processes of asexual embryogenesis, both in vivo and

in vitro, often differ from the regular divisions and patterning

events that deﬁ ne zygotic embryo genesis . Embryo pattern forma-

tion during apomictic embryogenesis ( parthenogenesis ) can be

irregular compared to zygotic embryogenesis in related sexual spe-

cies. In aposporous Hieracium , for example, embryogenesis fre-

quently commences earlier than in sexual plants as once the egg

differentiates, it transits rapidly to embryogenesis, and in some

cases altered division planes can result in a different embryo appear-

ance. Multiple embryos can also form in aposporous Hieracium

embryos in either the same or a secondary embryo sac [

58 ].

Although most Hieracium parthenogenetic embryos resemble

those formed by zygotic embryogenesis in sexual plants, embryos

with one or three cotyledons have also been observed. Despite

developmental alterations in the primary pattern of embryos

formed in aposporous Hieracium species, the resulting germinated

seedlings eventually exhibit normal plant growth when grown on

hormone free media in vitro [

58 ].

In vivo asexual embryogenesis in Kalanchoë species proceeds

through the typical globular, heart and torpedo stages from meri-

stematic cells along leaf margins [

24 ]. However unlike zygotic

embryo s, Kalanchoë asexual plantlets resemble shoots that then

grow adventitious roots from a hypocotyl structure [

23 ]. Once the

root system has developed, Kalanchoë plantlets detach from the

mother plant, fall to the ground and become new plants.

Melanie L. Hand et al.

11

In vitro embryogenesis could also be described as heteroge-

neous, as multiple developmental pathways are possible which

occur at varying frequencies within a single species and even the

same culture [

33 , 59 , 60 ]. Detailed characterization of in vitro

embryogenesis pathways has been performed using time-lapse

tracking from embryonic cell suspension s [

33 , 61 ]. Early

development of most microspore derived embryos involves a glob-

ular embryo with little cellular organization that undergoes sym-

metrical division and does not resemble a typical zygotic embryo

[

54 ]. Other microspore-derived embryos appear to form via a

developmental pathway that involves asymmetric division and con-

sequently more closely resemble zygotic embryos. Recently, micro-

spore embryogenesis systems have been developed that consistently

produce such embryos [

59 , 62 ]. These systems involve a heat stress

period that is either shorter or at a much lower temperature than is

usually applied.

Early during zygotic embryo genesis , a region of the embryo

differentiates to become a suspensor that functions to connect the

embryo to surrounding tissues, thereby positioning the embryo

inside the seed [

63 ]. The suspensor also acts to transport nutrients

and hormones to the embryo. When microspore embryogenesis

more closely mimics zygotic embryogenesis, a recognizable sus-

pensor is always present, which suggests the suspensor plays a role

in supporting early patterning events [

59 , 62 , 64 ]. A suspensor is

also formed during in vivo asexual embryogenesis, although

throughout Citrus nucellar embryony, the suspensor becomes evi-

dent at a much later stage of development than in zygotic embryos

[

15 ]. In aposporous Hieracium , embryos that develop in the

micropylar end of the embryo sac always form a suspensor and

embryos that develop within secondary chalazal embryo sacs may

or may not form a suspensor and often arrest at the globular stage

[

65 ]. The development of suspensors in asexual embryogenesis

suggests that fertilization is not required for formation of the

suspensor.

Unlike asexual embryos formed in apomictic seeds which

undergo desiccation and dormancy as part of seed maturation,

embryos formed in vitro and in vivo in Kalanchoë develop directly

into seedlings. Despite not developing within a seed, in vitro

somatic embryo s also undergo some form of maturation and accu-

mulate late embryogenesis abundant (LEA) proteins, although

sometimes treatment with ABA is ﬁ rst required to induce matura-

tion [

66 ]. In vitro somatic embryos also accumulate seed storage

proteins , which are recognized as important for the future devel-

opment of in vitro somatic embryos into plants. Only those

embryos that have accumulated enough storage proteins and have

acquired desiccation tolerance will develop into normal plants

[

60 ]. A comparison between asexual in vivo and somatic in vitro

embryogenesis processes was performed by measuring the

In Vitro and In Vivo Asexual Embryogenesis

12

accumulation of citrin seed storage proteins in polyembryonic

seeds and in vitro cultured embryos in Citrus . This study revealed

that in vitro embryos accumulate fewer citrins and at a later devel-

opmental stage than within the polyembryonic seed, suggesting

that despite not being derived from fertilization events, the nucel-

lar embryos are inﬂ uenced by the seed environment [

19 ].

Formation of endosperm is a crucial component of seed devel-

opment which does not accompany in vitro embryogenesis. The

precursor of the endosperm is the large diploid central cell of the

embryo sac. During sexual seed formation, the endosperm will

only develop following double fertilization , when one of the two

sperm cells fuses with the two central cell nuclei to produce trip-

loid endosperm. Formation of viable seed via apomixis also

requires the formation of endosperm. The majority of apomictic

species studied require fertilization to develop endosperm, a pro-

cess which is termed pseudogamy. In some apomictic species, typi-

cally members of the Asteraceae, endosperm can develop without

fertilization of the central cell. Maternal (m) and paternal (p)

genome ratios in the endosperm are typically 2m:1p in sexual spe-

cies and disturbance in this ratio may lead to seed abortion.

Apomicts tend to tolerate variation in endosperm ploidy and

maternal and paternal genome ratios which are not easily tolerated

in sexually reproducing plants, and have developed various strate-

gies to ensure seed viability [

67 ].

Apomictic Hieracium species are able to form endosperm

without fertilization . The polar nuclei fuse prior to the develop-

ment of nuclear and then cellular endosperm, in the absence of

fertilization and the resulting endosperm exhibits a 4m:0p genome

ratio in aposporous Hieracium . The trait of autonomous endo-

sperm (AutE) has recently been separated from fertilization-

independent embryogenesis in Hieracium through two

inter-speciﬁ c crosses [

68 ]. Two individuals were identiﬁ ed that

form reduced embryo sacs containing meiotically derived eggs and

central cells through the sexual pathway. However, egg cell s within

these individuals are unable to commence embryogenesis without

fertilization although in the absence of fertilization, the fused polar

nuclei undergo proliferation and continue to develop cellular

endosperm with a 2m:0p genome ratio. This indicates a paternal

genome contribution is neither required for endosperm initiation,

nor cellularization in both chromosomally reduced and unreduced

embryo sacs. When egg cells from these individuals are fertilized,

embryogenesis occurs to completion and viable seed is formed. It

is currently unclear if the central cell is also able to be fertilized as

this would result in a parental genome ratio of 2m:1p ratio as seen

in sexual species [

68 ].

Melanie L. Hand et al.

13

5 Genes Implicated in In Vivo and In Vitro Asexual Embryogenesis

Similarities between asexual embryogenesis in vitro and in vivo

raise questions regarding whether these processes are controlled by

the same molecular mechanisms. Although no genes responsible

for embryogenesis have yet been isolated from apomictic plants, a

number of gene candidates have been identiﬁ ed through differen-

tial gene expression analysis, genetic mapping and study of sexual

mutants with phenotypes that mimic asexual embryogenesis.

Attempts to understand in vitro somatic and gametic embryogen-

esis have also resulted in a range of gene candidates that when

expressed ectopically, result in embryo formation.

One of the ﬁ rst genes associated with somatic embryo genesis

was SOMATIC EMBRYOGENESIS RECEPTOR KINASE

( SERK ) when its involvement was demonstrated in carrot cell cul-

tures [

69 ]. SERK was identiﬁ ed as a marker for cells transitioning

from a somatic to an embryogenic state, due to its transient expres-

sion in established suspension cell cultures [

69 ]. SERK is a leucine-

rich repeat (LRR) receptor-like kinase that is also expressed in

developing ovules and embryos in planta and may therefore inﬂ u-

ence somatic embryogenesis through the same mechanisms of the

sexual pathway [

69 , 70 ]. Overexpression and downregulation of

SERK increases and decreases the efﬁ ciency of somatic embryo-

genesis, respectively [

70 , 71 ]. Interestingly, a SERK gene has also

been implicated in asexual reproduction within an apomictic grass,

Poa pratensis . cDNA-AFLPs differentially expressed between apo-

mictic and sexual lines of P. pratensis revealed a SERK gene that

displays differential expression [

72 ]. Apomixis in P. pratensis

involves development of an embryo sac not from the megaspore

mother cell (MMC) which is the typical precursor cell for the sex-

ual pathway, but from a diploid somatic cell positioned nearby the

MMC. These somatic precursor cells are found in the nucellar

ovule tissue. Within P. pratensis , SERK is expressed in embryo sac

precursor cells: the MMC in sexual plants, and somatic nucellar

cells in apomictic plants [

72 ]. The same expression proﬁ le was also

observed in apomictic and sexual lines of Paspalum notatum [

73 ].

SERK expression was also examined in apomictic Hieracium where

it was detected throughout the ovule, and expression was not

restricted to the nucellar region or MMC in Hieracium. SERK

expression was also observed in developing Hieracium embryos

[

74 ]. SERK is therefore thought to play an important role in

changing developmental fate of cells, both in stages of apomixis

and in somatic embryogenesis. BABYBOOM ( BBM ) is another

gene that has been associated with both in vitro and in vivo asexual

embryogenesis. BBM is an APETELA2 (AP2) transcription factor

that was originally identiﬁ ed following subtractive hybridization of

cDNA from Brassica napus microspores undergoing

In Vitro and In Vivo Asexual Embryogenesis

14

embryogenesis [ 75 ]. Ectopic expression of BBM in Arabidopsis or

B. napus induces somatic embryos, and constitutive expression of

BBM gene s from other species also results in the emergence of

ectopic embryos [

75 – 77 ]. BBM expression was also observed in

developing Arabidopsis zygotic embryo s [

75 ]. These results sug-

gest that BBM has a conserved role in the induction and/or main-

tenance of embryo development . BBM genes have also been

identiﬁ ed within a genomic region essential for apomixis in the

apomictic grass Pennisetum squamulatum [

78 ]. The apospory-

speciﬁ c genomic region (ASGR) of Pennisetum was identiﬁ ed fol-

lowing marker analysis of a selection of apomictic and sexual plants,

which revealed a set of apomixis-speciﬁ c markers that deﬁ ne the

ASGR [

79 ]. Sequencing of BAC clones from within the ASGR

revealed putative protein coding regions, including two of which

had similarity to BBM of rice [

78 ]. The ASGR is thought to con-

tain genetic elements responsible for both the formation of a dip-

loid embryo sac, and the process of parthenogenesis . The BBM

genes within the ASGR are therefore candidate apomixis genes

with strong potential to have a role in the induction or mainte-

nance of asexual embryogenesis in Pennisetum apomicts. However,

conﬁ rmation of a role for BBM in parthenogenesis has not yet been

reported.

The involvement of common genes in zygotic and asexual

embryogenesis implies that despite arising from different activa-

tion signals and different tissues, each embryogenesis process con-

verges on a similar developmental pathway. Genes with a known

involvement in zygotic embryo genesis have therefore been studied

in asexual embryogenesis systems to understand whether such

genes are also involved in asexual embryogenesis. The LEAFY

COTYLEDON ( LEC ) family of transcription factor s is crucial for

regular embryogenesis and is also implicated in somatic embryo-

genesis. Arabidopsis contains three LEC gene s : LEC1 , LEC2, and

FUSCA3 ( FUS3 ), and each of these genes is expressed exclusively

in the embryo [

80 – 82 ]. Ectopic expression of each of the three

LEC genes leads to vegetative cells adopting characteristics of

maturation- phase embryos, and hence this gene family is associ-

ated with the process of somatic embryogenesis [

80 – 82 ]. The LEC

genes have been linked to auxin production, as LEC2 is known to

activate the auxin biosynthesis genes YUCCA2 and YUCCA4

[

83 ]. FUS3 expression also increases in response to auxin [ 84 ].

This interaction with auxin signaling is thought to be responsible

for the ability of LEC gene expression to induce embryonic

competence.

LEC1 has been studied in Kalanchoë species and is implicated

in the process of asexual plantlet formation in these species.

Compared to Arabidopsis , the LEC1 gene of Kalanchoë daigre-

montiana ( KdLEC1 ) is truncated and does not rescue the

Arabidopsis lec1 mutation, suggesting it functions differently to

Melanie L. Hand et al.

15

LEC1 in Arabidopsis [ 23 ]. A functional full length copy of LEC1

was created by replacing the deleted nucleotides in KdLEC1 with

the corresponding nucleotides from Arabidopsis and transforma-

tion of Kalanchoë daigremontiana with this synthesized LEC1-

LIKE gene results in disrupted asexual reproduction and in some

instances abortion or absence of plantlet formation [

85 ]. This

study strongly supports the involvement of LEC1 in in vivo asexual

embryogenesis in Kalanchoë and furthermore suggests that the

switch from sexual to asexual propagation in the evolution of

Kalanchoë was probably activated following truncation of the

KdLEC1 gene [

85 ].

Another gene that appears to be involved in the induction of

an embryogenic state is the RWP-RK domain containing (RKD)

transcription factor RKD2, which is preferentially expressed in the

egg cell of Arabidopsis and wheat [

86 ]. Ectopic expression of

RKD2 results in ovule integument cells that become enlarged and

densely cytoplasmic with prominent nuclei, suggesting these cells

have become pluripotent [

87 ]. Ectopic RKD2 expression also

results in some integument cells adopting an egg cell identity, and

a low frequency (ca. 0.1 %) of embryo-like structures also appear

outside of the embryo sac [

87 ]. This observation is reminiscent of

adventitious embryony and may indicate that RKD2 is involved in

the induction of embryogenesis from ovule tissue during adventi-

tious embryony.

Additional genes including WUSCHEL and AGAMOUS-Like

15 ( AGL15 ) are known to induce embryo formation from vegeta-

tive tissue when ectopically expressed, and have therefore been

implicated in somatic embryo genesis [

88 , 89 ]. WUSCHEL is

known to be involved in specifying and maintaining stem cells in

the shoot and root meristem [

90 ] while AGL15 is known to accu-

mulate in developing embryos [

91 ], therefore a role in embryo-

genesis is to be expected for both of these genes. However, with

the exception of SERK , most of the genes shown to be involved in

zygotic and asexual embryogenesis are not speciﬁ cally expressed in

the egg cell or the zygote. Therefore, whilst important for later

stages of embryo development , these genes may not be involved in

the process of embryo initiation which is possibly the most impor-

tant aspect of asexual embryogenesis. It has been proposed that the

observed ectopic embryo development associated with mis-

expression of these genes, is a result of cellular stress, rather than a

speciﬁ c initiation signal expressed by the genes [

92 ]. This hypoth-

esis is consistent with embryonic competence being induced by

stress factors, as discussed earlier.

To understand the genetic elements responsible for inducing

embryonic competence in both in vitro and in vivo asexual embryo-

genesis, future experiments will likely focus on comparison of gene

expression from embryo precursor cells directly before and after

the initiation of embryogenesis. Genetic mapping of apomixis loci

In Vitro and In Vivo Asexual Embryogenesis

16

may also reveal which genes are responsible for the initiation of

asexual embryogenesis. Genetic analyses of apomicts have shown

that gametophytic apomixis is inherited as a dominant trait. In

many apomictic species, developmental components of apomixis

(meiotic avoidance and parthenogenesis ) are controlled by inde-

pendent loci and further research is underway to isolate the causal

sequences that underlie these loci. For example, characterized

deletion mutants developed in apomictic Hieracium praealtum

revealed a genomic region responsible for fertilization- independent

embryogenesis and endosperm formation, named LOSS OF

PARTHENOGENESIS ( LOP ) [

93 ]. Deletion of LOP sees the

plant become dependent upon fertilization for both embryo and

endosperm development (Fig.

2 ) [ 13 ]. Genetic mapping of LOP

and AutE is the focus of current work that may lead to isolation of

the causal sequences for both traits.

A genomic locus strongly associated with adventitious embry-

ony in Citrus has also been identiﬁ ed [

94 ]. Further characteriza-

tion of this locus may clarify the mechanism of adventitious

embryony and identify the genetic element responsible for

Fig. 2 Cleared ovules from wildtype apomict Hieracium praealtum ( a ), and a H.

praealtum lop mutant ( b ) that has lost the capacity to undergo parthenogenesis

and autonomous endosperm development. Within apomictic H. praealtum ovules,

embryo and endosperm develop from the egg and central cell, respectively, with-

out fertilization ( a ). H. praealtum lop deletion mutant m179 ( b ) has lost this

capacity and the egg and central cells do not develop further without fertilization.

Scale bars = 50 μm. em embryo, e egg, cc central cell, ne nuclear endosperm.

Ovules were collected at stages 6 ( a ) and 10 ( b ) of capitulum development

according to Koltunow et al. [ 65 ]

Melanie L. Hand et al.

17

inducing asexual embryogenesis in planta . Similarly, the search for

genes within the controlling parthenogenesis loci of the salmon

wheat system may reveal those genes that are responsible for par-

thenogenesis in this system. Although controlling genes are cur-

rently unknown for the salmon wheat system, it is likely that they

have lost those genes required for repressing fertilization indepen-

dent embryogenesis in sexual plant species.

6 Epigenetic Inﬂ uence on Asexual Embryogenesis

Various epigenetic marks and pathways have been associated with

both sexual and asexual embryogenesis processes, suggesting that

the induction and regulation of asexual embryogenesis may involve

epigenetic components. For instance, the application of exogenous

auxin during somatic embryo induction results in DNA hyper-

methylation [

95 ], and inhibition of DNA methylation suppresses

the formation of embryogenic cells from cultured carrot epidermal

cells [

96 ]. Auxin could therefore possibly reprogram gene expres-

sion through DNA methylation, leading to the induction of

embryogenesis pathways within somatic cells.

Genes within epigenetic pathways have also been implicated in

both in vitro and in vivo asexual embryogenesis. One such epigen-

etic factor implicated in asexual embryogenesis is PICKLE (PKL),

a chromatin remodeling protein [

97 ]. PKL is responsible for

repressing the LEC family of transcription factor s, as pkl mutants

display overexpression of LEC1 , LEC2 and FUS3, and display a

phenotype similar to that seen from LEC1 overexpression [

97 ,

98 ]. PKL activity is therefore acknowledged as an important regu-

latory mechanism for repressing embryonic identity throughout

seedling growth, by suppressing the embryogenic program in

somatic cells [

99 ]. For this reason, PKL is also a candidate for the

induction of asexual embryogenesis. Deregulation of PKL in

somatic cells or within the egg cell would permit expression of

embryogenic genes that are generally only expressed by the devel-

oping embryo following fertilization . However to date, PKL has

not been speciﬁ cally associated with asexual embryogenesis in any

natural apomictic plant.

Strong evidence exists suggesting that epigenetic pathways

play a crucial role in asexual embryo and endosperm development

during apomixis . Mutants of the Polycomb -Group (PcG) chroma-

tin modeling complex show phenotypes reminiscent of fertilization

independent embryogenesis and endosperm formation seen in

gametophytic apomixis. In particular, the Polycomb Repressive

Complex 2 (PRC2) is known to be involved in the suppression of

seed development in the absence of fertilization. The PRC2 is con-

served between plants and animals and represses gene expression

via trimethylation of histone H3 at lysine 27 (H3K27me3).

In Vitro and In Vivo Asexual Embryogenesis

18

Phenotypes of asexual embryo and endosperm development have

been observed when core PRC2 genes are mutated in Arabidopsis .

For instance, the fertilization-independent seed (FIS) PRC2

complex (FIS-PRC2) consists of the genes MEDEA ( MEA ), FIS2 ,

FERTILIZATION-INDEPENDENT ENDOSPERM ( FIE ), and

MULTICOPY SUPPRESSOR OF IRA1 ( MSI1 ). Loss of function

of any of these genes results in endosperm initiation and prolifera-

tion without fertilization. However, the endosperm does not cel-

luarize [

100 – 102 ]. The role of the FIS-PRC2 complex is therefore

considered to inhibit central cell proliferation. In the case of MSI1

mutants, low levels of parthenogenetic embryo initiation are

observed, followed by embryo arrest, so that viable seeds are not

formed [

103 ]. The role of some of the FIS-PRC2 genes has been

investigated during seed initiation in Hieracium spp., one of the

few groups of apomicts that develop endosperm without fertiliza-

tion. Downregulation of Hieracium FIE (HFIE), a protein linking

multiple PRC2 components inhibiting fertilization-independent

endosperm proliferation in Arabidopsis does not result in

fertilization- independent endosperm proliferation in sexual plants.

HFIE function is required for completion of both sexual and asex-

ual embryo and endosperm development in examined Hieracium

species [

104 ]. These results demonstrate that the capacity for

embryogenic competence and endosperm formation in apomicts

may function via deregulation of other PRC2 complex family

members and that additional factors are required to produce viable

asexual embryos and endosperm. The identiﬁ ed Hieracium AutE

plants that form endosperm, but not embryos, without fertiliza-

tion may help identify and deﬁ ne the roles of genes that regulate

the autonomous endosperm mechanism.

The PRC2 complex interacts with other genes implicated in

asexual embryogenesis, including the LEC gene family. LEC1 ,

LEC2, and FUS3 are all overexpressed in CURLY LEAF ( CLF ) and

SWINGER ( SWN ) double mutants, which are PcG gene homo-

logues of the PRC2 gene MEA [

105 ]. A cis regulatory element has

been identiﬁ ed within the LEC2 promoter which is responsible for

recruiting the PRC2 complex [

106 ]. These results suggest that the

PcG acts to repress embryonic gene expression by histone methyla-

tion. Histone acetylation is another epigenetic mark that in contrast

to histone methylation, is generally associated with transcriptional

activation. Removal of the acetylation is performed by histone

deacetylase (HDAC), which consequently results in transcriptional

repression. Interestingly, two histone deacetylase genes ( HDA6 and

HDA19 ) are partly responsible for repressing the embryonic pro-

gram during Arabidopsis germination [

107 ]. Another HDAC gene

( HDA7 ) in Arabidopsis is known to be important for normal

embryo development [

108 ]. Inefﬁ cient or defective histone deacet-

ylation of key embryonic genes may therefore be a candidate mech-

anism for inducing asexual embryogenesis.

Melanie L. Hand et al.

19

7 Conclusions

While the pathways involved in developing the embryo itself appear

common between the various modes of embryogenesis described

here, many differences exist between the initiation processes of

asexual embryogenesis in vitro and in vivo. Unlike somatic or

gametic embryogenesis , apomixis -associated embryogenesis occurs

near maternal reproductive tissue, and develops within a seed

structure. Despite these differences, in vitro and in vivo asexual

embryogenesis share some common factors: a change in the devel-

opmental fate of embryogenic precursor cells; and expression of an

embryonic pathway in such cells without fertilization . Identifying

molecular mechanisms that underlie these processes within in vitro

systems may help to understand pathways that lead to apomixis.

While some candidate genes for both in vitro and in vivo asexual

embryogenesis have been identiﬁ ed, a role in apomixis has not yet

been conﬁ rmed for any of these genes. One possibility is that

embryogenesis related genes are deregulated by epigenetic factors

during asexual embryogenesis. Continued research into asexual

embryogenesis will yield important ﬁ ndings related to plant cell

fate speciﬁ cation and the molecular regulation of embryogenesis.

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In Vitro and In Vivo Asexual Embryogenesis

25

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_2, © Springer Science+Business Media New York 2016

Chapter 2

Somatic Versus Zygotic Embryogenesis: Learning

from Seeds

Traud Winkelmann

Abstract

Plant embryogenesis is a fascinating developmental program that is very successfully established in nature

in seeds. In case of in vitro somatic embryogenesis this process is subjected to several limitations such as

asynchronous differentiation and further development of somatic embryos, malformations and disturbed

polarity, precocious germination, lack of maturity, early loss of embryogenic potential, and strong geno-

typic differences in the regeneration efﬁ ciency. Several studies have shown the similarity of somatic and

zygotic embryos in terms of morphological, histological, biochemical, and physiological aspects. However,

pronounced differences have also been reported and refer to much higher stress levels, less accumulation

of storage compounds and a missing distinction of differentiation and germination by a quiescent phase in

somatic embryos. Here, an overview on recent literature describing both embryogenesis pathways, com-

paring somatic and zygotic embryos and analyzing the role of the endosperm is presented. By taking

zygotic embryos as the reference and learning from the situation in seeds, somatic embryogenesis can be

improved and optimized in order to make use of the enormous potential this regeneration pathway offers

for plant propagation and breeding.

Key words Biochemistry , Comparative approach , Maturation , Morphology , Proteome , Storage

reserves , Stress response , Transcriptome

1 Introduction

Somatic embryo genesis, a fascinating developmental pathway

through which plants can be regenerated from bipolar structures

derived from a single or a few somatic cells was ﬁ rst described more

than 50 years ago in carrot by Reinert [

1 ] and Steward et al. [ 2 ].

This regeneration pathway offers a great potential to be applied in

mass propagation, genetic transformation by direct means or via

Agrobacterium tumefaciens and as a source of protoplasts as well as

for long-term storage of germplasm using cryopreservation. Also

fundamental studies of early embryogenesis are easier to be per-

formed with somatic than with zygotic embryo s. However, up to

now the exploitation of this pathway is limited by inherent

26

problems that are observed in many different plant species, like

asynchronous differentiation and further development of somatic

embryo s, malformations and disturbed polarity , precocious germi-

nation , early loss of embryogenic potential , and strong genotypic

differences in the regeneration efﬁ ciency. On the other hand, such

limitations are not found in zygotic embryos developing within

seeds. Thus, this review aims at comparing these two types of

embryogenesis by regarding zygotic embryogenesis as a reference

as suggested for the ﬁ rst time for wheat by Carman [

3 ]. The iden-

tiﬁ cation of the major differences could enable new approaches to

optimize somatic embryogenesis. Available literature dealing with

comparisons of somatic and zygotic embryos on morphological,

histological, biochemical, and also transcriptomic and proteomic

level will be summarized, with emphasis on our model plant, the

ornamental species Cyclamen persicum .

The zygote is formed after double fertilization has taken place

which is leading to the formation of the embryo and the endo-

sperm . Zygotic embryo genesis is a complex, highly organized pro-

cess, that has been studied for a long time by histological approaches

only [

4 ]. Recently it has been supplemented by molecular genetic

studies, mainly based on mutant analyses of Arabidopsis thaliana as

excellently reviewed in 2013 by Wendrich and Weijers [

5 ] and

depicted in Fig.

1 . Embryogenesis is divided into (1) embryogenesis

sensu strictu (morphogenesis of embryo and endosperm) meaning

the development of the zygote up to a cotyledonary stage embryo

and (2) the subsequent maturation phase that starts with the switch

from maternal to ﬁ lial control [

6 ] and ﬁ nally (3) the phase of

embryo growth and seed ﬁ lling ending with a desiccation phase [

7 ].

Embryogenesis sensu strictu starts with a loss of polarity directly

after fertilization of the egg cell which is followed by re- polarization

and elongation of the zygote [

5 ] . The important ﬁ rst asymmetric

cell division of the zygote results in a more elongated basal cell that

gives rise to the suspensor and the hypophysis and a small apical

cell that generates the embryo. The suspensor positions the embryo

within the embryo sac, conducts nutrients to the developing

embryo and is a source of plant hormones that are important for

polarity establishment [

8 ]. It is eliminated by programmed cell

death between globular and torpedo stage in angiosperms and in

late embryogenesis in gymnosperms [

9 ].

Auxin is the predominant plant hormone that has been

reported to be involved in polarity and pattern formation.

Especially, the PIN (PIN formed proteins) dependent asymmetric

auxin efﬂ ux regulates these processes in early embryogenesis ( [

10 ] ,

reviewed in 2010 by De Smet et al., [

11 ]). The role of other plant

hormones, among which cytokinins and brassinosteroids were

reported to be important in these processes, is not yet clearly

resolved [

11 ].

1.1 Zygotic

Embryogenesis

Traud Winkelmann

27

Subsequent organized cell division in a symmetric way and

only in one direction leads to the formation of the suspensor . In

the apical cell division planes change in a strictly regulated way in

A. thaliana and thereby establish two types of axes, deﬁ ning upper

and lower tiers and radially arranged cell types [

5 ]. Most interest-

ingly, the ﬁ rst cell divisions take place within the space provided by

the apical cell. Thus, pattern formation occurs in the globular

embryo by which the protoderm cells, vascular and ground tissue

are deﬁ ned. The last stage of embryogenesis sensu strictu is the

heart stage being characterized by the presence of shoot and root

apical meristems as well as early cotyledons. The key genes regulat-

ing morphogenesis of the embryo have been identiﬁ ed and encode

transcription factor s, receptor kinases, proteins involved in plant

hormone signaling and micro RNAs pointing to the predominant

Fig. 1 Morphogenetic processes during Arabidopsis embryogenesis. Schematic overview of Arabidopsis

embryogenesis from the egg cell to the heart stage embryo, highlighting the morphogenetic processes required

to progress from one stage to the next. The colors represent cells of (essentially) the same type ( see color

legend), based on marker gene expression and lineage analysis. Cot cotyledon, SAM shoot apical meristem,

Hyp hypocotyl, RAM root apical meristem (reproduced from [ 5 ] with permission from New Phytologist)

Somatic Versus Zygotic Embryogenesis

28

transcriptional control, and future research needs to focus on how

these regulators hold their function in terms of cell biological

implementations [

5 ] .

The later phase of seed development (maturation phase) com-

prises embryo growth, seed ﬁ lling by deposition of storage reserves

and ﬁ nally desiccation . Mainly seed dormancy has attracted the

attention of research in A. thaliana and other species (reviewed by

Finch-Savage and Leubner-Metzger in 2006, [

12 ]). Seed ﬁ lling is

of importance for many agricultural crops like rape seed or legumes

as well (reviewed by Verdier and Thompson in 2008, [

13 ]). At the

end of seed development, the zygotic embryo is in a quiescent

state which clearly separates embryogenesis from germination .

The term somatic embryo genesis already points to the pronounced

morphological similarity of this vegetative regeneration pathway to

zygotic embryo genesis . Somatic embryo genesis generally starts

from a single cell or a group of cells of somatic origin and direct

somatic embryogenesis is distinguished from indirect somatic

embryogenesis in which a callus phase is passed through. The

induction of embryogenic cells sometimes refers to all events that

reprogram a differentiated cell into an embryogenic cell, but

recently was divided into different phases, i.e., dedifferentiation,

acquisition of totipotency , and commitment into embryogenic

cells [

14 ]. The ﬁ rst important difference compared to zygotic

embryogenesis is the need for both, transcriptional and transla-

tional reprogramming of a somatic cell. Dedifferentiation of the

somatic cells is the prerequisite to gain embryogenic competence

and results in genetic reprogramming, loss of fate, and change into

meristematic cells [

15 ]. Stress due to wounding, separation from

surrounding tissue, in vitro culture conditions, and also auxin are

discussed to have a pivotal role in dedifferentiation [

15 ]. Elhiti

et al. [

14 ] postulated that cells have to be cytologically separated

for dedifferentiation as expression of genes responsible for second-

ary cell wall formation changed. Moreover, pronounced changes in

the network that regulates the response to hormones have to take

place. Twenty-ﬁ ve candidate genes being associated with the

expression of cellular totipotency were identiﬁ ed by a bioinfor-

matic approach using the CCSB (Center of Cancer Systems

Biology) interactome database and Arabidopsis as a model for a

molecular regulation network [

14 ]. They cover functions in tran-

scription, signal transduction, posttranslational modiﬁ cation,

response to plant hormones, DNA repair and DNA methylation,

and for the ﬁ rst time protein phosphorylation and salicylic acid

signaling. The ﬁ nal step of the induction phase, the commitment

into embryogenic cells, involves genes for signal transduction,

microtubule organization, DNA methylation, regulation of tran-

scription, apoptosis, and hormone-mediated signaling [

14 ].

1.2 Somatic

Embryogenesis

Traud Winkelmann

29

The establishment of polarity and a ﬁ rst asymmetric cell divi-

sion has been observed in early somatic embryo genesis of carrot

[

16 ] and alfalfa [ 17 ]. By cell tracking experiments it was shown

that carrot somatic embryos developed from different single sus-

pension cells either via a symmetric or via an asymmetric ﬁ rst divi-

sion [

18 ], indicating that an asymmetric division is not decisive for

proper somatic embryo development . However, as stated by Feher

et al. [

15 ] , polarity, in terms of the transcriptional and biochemical

status of the cell, is not necessarily expressed at the level of the

morphology and symmetry of cell division. Therefore, early polar-

ization is thought to be crucial in somatic embryogenesis as well as

in zygotic embryo genesis , but needs to be set up by the cell inter-

nally following an external stimulus. The suspensor originating

already from the ﬁ rst asymmetric division of the zygote is also

formed in somatic embryos of conifers. It is supposed to support

polarity and axis establishment in embryos and undergoes pro-

grammed cell death also in somatic embryos (reviewed by

Smertenko and Bozhkov in 2014, [

8 ]). In contrast, suspensor

structures are often not so clearly detectable or completely missing

in somatic embryos of plant species other than gymnosperms .

Due to the difﬁ culty of identiﬁ cation of embryogenic cells, the

early stages up to the globular embryo, and especially the precise

sequence of cell divisions that can be described for Arabidopsis zygotic

embryo genesis resulting in pattern formation have not often been

recorded in somatic embryo genesis systems. Most studies that

track the development of somatic embryos start with the globular

stage [

4 ]. Further development runs through the typical stages of

angiosperm embryogenesis in dicots, namely globular stage, heart

stage, torpedo stage, and cotyledonary stage. For a long time,

markers for competent cells have been searched for, and most

promising are Somatic Embryogenesis Receptor like Kinases

(SERKs), that were identiﬁ ed to play a role in zygotic and somatic

embryogenesis in Daucus carota [

19 ] and A. thaliana [ 20 ]. They

are involved in perception and transduction of extracellular signals

and connected to brassinosteroid signaling [

21 ], but their exact

function is unknown up to now.

Maturation includes accumulation of storage reserves , growth

arrest, and acquisition of desiccation tolerance and is, in case of

somatic embryo s, induced externally by increasing the osmotic

pressure (lowering the osmotic potential) of the culture media

(e.g. by addition of polyethylene glycol or increased sugar concen-

tration) and application of abscisic acid ( ABA ) [

22 ]. Germination

requires similar conditions as in the respective zygotic embryo s

and completes this developmental pathway. Obviously, somatic

embryos are completely lacking the effects of the surrounding seed

tissues which provide physical (space) constraints and a speciﬁ c and

complex interaction of testa and endosperm supporting embryo-

genesis in an optimal way. For the induction of embryogenic cells,

Somatic Versus Zygotic Embryogenesis

30

external stimuli are mainly coming from the culture media, plant

growth regulator s , and culture conditions, but thereafter somatic

embryogenesis is following an intrinsic autoregulatory develop-

mental program [

8 ]. Most likely, this process can be improved by

mimicking conditions found in seeds.

2 Comparison of Somatic and Zygotic Embryos

The fact that somatic embryo genesis was named after embryogen-

esis taking place in seeds clearly indicates a high degree of similarity

of somatic and zygotic embryo s. Many early studies were devoted

to describe morphological aspects involving histological and micro-

scopic investigations. Due to the typical stages both types of

embryos pass through, globular, heart, torpedo, and cotyledonary

stage, the parallels become obvious. Both kinds of embryos are

bipolar structures from the beginning and do not have a vascular

connection to maternal tissue which enables the discrimination of

somatic embryogenesis and adventitious shoot regeneration.

The ﬁ rst cell division of the zygote is asymmetric while in

somatic embryo s this is not always the case (see above, [

18 ]).

Mathew and Philip [

23 ] described the regeneration of Ensete super-

bum via somatic embryogenesis starting from single cells without

the need of strong polarity establishment in these cells. However,

all further stages that were compared in this histological approach

revealed high similarity of somatic embryos to their zygotic coun-

terparts in terms of structure of the embryonic apex or formation

of cotyledons and hypocotyls. In many indirect somatic embryo-

genesis systems, the so-called proembryogenic masses, being clus-

ters of small, dense cytoplasm rich embryogenic cells, give rise to

the differentiating embryos, but their ﬁ rst divisions have not often

been observed in detail, since the cell or the cell group from which

the embryo originates is difﬁ cult to identify. While in gymnosperm

somatic embryos the suspensor is a very prominent structure that

in late embryogenesis undergoes programmed cell death [

8 ], in

many angiosperm systems suspensors are either absent or strongly

reduced which might explain the difﬁ culties in root formation

reported for some species, especially due to the absence of the

hypophysal cell [

4 ].

Maize secondary somatic embryo s derived from single primary

somatic embryos or somatic embryos developing attached to callus

cells, revealed malformations in the shoot meristem formation after

direct regeneration of the single somatic embryos, while those that

developed next to callus cells perfectly represented zygotic embryo

development [

24 ]. The authors discuss a possible role of the neigh-

boring callus cells with similar functions as suspensor cells in the

zygotic situation. Interestingly, in our model plant C. persicum

2.1 Morphological

and Histological

Comparison

Traud Winkelmann

31

[ 25 ] embryogenic cultures are mixtures of embryogenic and

nonembryogenic cells, and the differentiating somatic embryos are

surrounded by a extracellular matrix resembling several cell wall

layers (Douglas Steinmacher, Melanie Bartsch, and Traud

Winkelmann, unpublished data). One possible explanation, for

which further evidence is needed, could be that nonembryogenic

cells undergo programmed cell death and thereby enable differen-

tiation. In Eucalyptus nitens somatic embryos are only sporadically

observed, but then appear on dark brown wounded callus cells

[

26 ]. An ultrastructural study not only recorded several analogies

in cell and embryo structure when compared to zygotic embryos,

it also identiﬁ ed a kind of waxy coat surrounding the somatic

embryos which was supposed to originate from phenolic exudates

[

26 ]. Somatic embryo s of C. persicum have three times larger cells

than their zygotic counterparts, and their outer surface is more

irregular than the smooth protoderm of zygotic embryos [

27 ].

This observation indicates that the physical and chemical con-

straints of the surrounding tissue, the endosperm , may have an

important inﬂ uence on the cellular organization of zygotic embryos

that is lacking in somatic embryogenesis systems ( see also

Subheading

3 ).

Maturation is a major bottleneck in somatic embryo genesis of

several species including Pinus pinaster [

28 ] and coffee [ 29 ]. Also

loblolly pine somatic embryos did not reach full maturity and had

lower dry weights than the zygotic ones [

30 ]. Polyethylene glycol

( PEG ) which is often used in maturation media of conifers had

clear effects on the morphology of somatic embryos of P. pinaster

as numerous and larger vacuoles as well as larger intercellular spaces

were induced by this treatment [

28 ]. By the histological compari-

son of somatic embryos subjected to different maturation treat-

ments ( carbohydrates in various concentrations) protein bodies

were found to appear earlier in somatic embryos, and to be more

abundant in well-developed somatic embryos leading to the sug-

gestion that storage protein accumulation could be regarded as a

marker for embryo quality of Pinus pinaster [

28 ]. The same authors

observed starch accumulating in zygotic embryo s in a gradient of

higher concentrations at the basal end, whereas in somatic embryos

the localization of starch granules strongly depended on the matu-

ration treatment. However, irrespective of the maturation treat-

ment, somatic embryos always contained higher amounts of starch

than the zygotic ones again with signiﬁ cant differences between

different kinds and concentrations of carbohydrates applied [

28 ].

Another aspect, namely the water status, was studied in Hevea

brasiliensis embryos [

31 ]. In zygotic embryo s the water content

decreased sharply from 91 to 53 % within 1 week (14–15 weeks

after pollination) and during the remaining maturation phase down

to 42 %. In contrast, somatic embryo s without maturation treat-

ments had a water content of nearly 80 %, while those that had

Somatic Versus Zygotic Embryogenesis

32

been desiccated or cultivated on higher sucrose concentrations

plus ABA still contained 71 % water but had much higher germina-

tion and conversion rates than the nontreated ones [

31 ]. Also in

date palm the zygotic embryos underwent dehydration with a

water content of 80 % decreasing to 35 %, whereas somatic embryos

had a water content of around 90 % throughout the whole devel-

opment [

32 ]. Both mentioned species still have high water content

in the seed after desiccation . In species with true orthodox seeds

and much lower water contents, the drop in water content and

thereby the discrepancy between somatic and zygotic embryos can

be expected to be even more pronounced.

Somatic embryo genesis is already commercialized in coffee , but

its proﬁ tability is limited due to losses during conversion into plant-

lets. Thus, Etienne et al. [

29 ] put special emphasis on studying this

phase in the zygotic and somatic system. Differences were found in

conversion time which took 22 weeks in somatic and 15 weeks in

zygotic embryo s, hypocotyl length being shorter in somatic

embryo s, a more spongy tissue in the somatic embryo axis, earlier

differentiation of stomata in somatic embryos and less protein and

starch in cotyledonary somatic embryos [

29 ] . The water content of

zygotic embryos increased strongly during germination starting

from 28 % and reaching 80 % within 4 weeks, whereas the increase

in somatic embryos was rather mild (water content from 70 to 85

%). Furthermore, the authors observed asynchronous germination

in somatic embryos. It can be concluded that the phase of matura-

tion which includes a growth arrest controlled by plant hormones

(mainly ABA ) and desiccation is obviously extremely important to

allow the development of high quality somatic embryos that will

germinate in high rates and in a synchronized way.

When screening the literature for studies comparing somatic and

zygotic embryo s on the biochemical level, mainly analyses of major

storage compounds, i.e. storage proteins , carbohydrates , and lipids

are found. Depending on the type of seed in a respective species,

storage reserves may be found in the embryo itself and here mainly

in the cotyledons or in the endosperm . Early studies in Brassica

napus [

33 ] and cotton [ 34 ] have shown that somatic embryo s are

able to accumulate storage proteins, but in much lower amounts

(1/10 of that found in zygotic embryos in B. napus ) and in earlier

stages. In somatic embryos of alfalfa 7S globulin was dominant,

while in zygotic embryos 11S globulin and 2S albumin were more

abundant [

35 ]. The processing and subcellular localization of 7S

and 11S storage proteins in protein bodies was comparable in both

embryo types, while 2S albumin in somatic embryos was detected

in the cytoplasm, in contrast to zygotic embryos in which 2S albu-

mins were localized in protein bodies [

35 ]. Overall, also in alfalfa

lower amounts of storage proteins were determined in somatic

embryos, thus supporting the observations in B. napus and cotton.

Thijssen et al. [

36 ] visualized globulin (storage protein)

2.2 Biochemical

Comparison

2.2.1 Storage Proteins

Traud Winkelmann

33

accumulation by ﬂ uorescence labeled antibodies in somatic and

zygotic embryos of maize. Starting 10 days after pollination globu-

lins were detected in the scutellum ﬁ rst and later in leaf primordia

and roots. Lower amounts of intermediate globulin precursor pro-

teins were found early in development of somatic embryos while

mature globulins could be induced by a maturation treatment with

ABA [

36 ]. Date palm somatic embryos contained about 20 times

lower amounts of total protein than zygotic embryos, a different

protein composition, and were lacking glutelin, a storage protein

with the typical accumulation and hydrolysis pattern in zygotic

embryos [

32 ]. In agreement with these studies are the observations

in oil palm embryos in terms of earlier, but 80 times less production

of 7S globulins in somatic embryos compared to zygotic ones [

37 ].

A recent follow-up study [

38 ] reported on early mobilization of

storage proteins by proteases in somatic embryos, thus providing

further evidence that the clear differentiation of the developmental

phases of embryogenesis, maturation, and germination is lacking in

somatic embryos. Instead there is an overlap of all three programs,

since globulin synthesis still occurred during germination of somatic

embryos and cystein proteases were active in all phases of somatic

embryogenesis [

38 ]. In order to gain insights into glutamine

metabolism, a nitrogen compound that is important for embryo-

genesis, Perez-Rodriguez et al. [

39 ] found cytosolic glutamine syn-

thase 1a (GS1a) to be absent in zygotic, but present in somatic

embryos of P. pinaster and Pinus sylvestris indicating the onset of

precocious germination in late stages of somatic embryogenesis,

since this gene is a marker for chloroplast differentiation. GS1b

expression was detected in procambial tissues of both types of

embryos with the level of expression correlating to the quality of

somatic embryos [

39 ]. Arginase expression in somatic embryos

indicated that storage protein breakdown obviously started before

germination [

39 ]. Possibilities to improve storage protein accumu-

lation by ABA treatment were shown for example for cocoa somatic

embryos [

40 ] or by increasing sucrose concentrations in matura-

tion media for Pinus strobus [

41 ] and cyclamen [ 42 ].

Cotyledonary white spruce somatic embryo s accumulated more

starch, but less proteins and lipids than zygotic embryo s in the

same stage. This points to the fact that the conversion of starch

into the energy rich storage compounds lipids and proteins did not

take place in somatic embryos to the same extent [

43 ]. According

to this study, adjustment of in vitro culture conditions might be an

option to improve this conversion during embryo maturation .

Carbohydrates have important functions during plant develop-

ment and growth as energy sources but also for osmotic adjust-

ment, protein protection, and signaling molecules, and they have

been analyzed in comparative approaches during somatic and

zygotic embryogenesis . During maturation of cocoa zygotic

embryos ( Theobroma cacao ) storage proteins and starch

2.2.2 Carbohydrates

Somatic Versus Zygotic Embryogenesis

34

accumulate, dehydration takes place and monosaccharides and

sucrose decrease, while two oligosaccharides, rafﬁ nose and stachy-

ose, increase [

40 ]. In contrast, somatic embryos accumulated less

protein and starch as detected in histological studies and they had

higher levels of sucrose, xylose, and rhamnose [

40 ]. A shift in car-

bohydrate composition was observed in Norway spruce for both,

somatic and zygotic embryos, during later developmental stages

with decreasing total carbohydrates and a higher sucrose:hexose

ratio within time. However, only mature zygotic embryos con-

tained rafﬁ nose and stachyose which play a role in desiccation tol-

erance [

44 ]. After a maturation treatment with 3.75 % PEG 4000

the sucrose:hexose ratio in Norway spruce somatic embryos raised

signiﬁ cantly from 0.88 to 6 which resembled more the ratio of 9.7

found in zygotic embryos, all in the early cotyledonary stage [

45 ].

While in somatic embryos invertase and sucrose synthase were

found in high activity during the proliferation and early maturation

phase, invertase activity was low in developing zygotic embryos

and sucrose synthase was ﬁ rst observed in the cell layer surround-

ing early zygotic embryos and later inside the embryos. From this

the authors conclude that sucrose synthase plays an important role

in the transition of the embryo from a metabolic sink to a storage

sink [

45 ]. The sucrose distribution within the embryo which is

among other factors controlled by epidermal sucrose transporters

was suggested to trigger starch accumulation during the matura-

tion phase of Vicia faba zygotic embryos [

46 ].

In the fruit tree Acca sellowiana that is native to South Brazil,

total soluble carbohydrates per gram fresh mass were found to be

about twice as high in zygotic compared to somatic embryo s in the

globular, heart, and torpedo stage, although the principal compo-

sition was the same. Especially for sucrose , fructose, myo-inositol ,

and rafﬁ nose (in the later stages of embryogenesis) zygotic embryo s

showed higher contents, even though somatic embryos were cul-

tured in sucrose containing media. On the other hand starch con-

tents of torpedo and cotyledonary stage somatic embryos exceeded

those of their zygotic counterparts [

47 ]. Also in pea changes in

soluble sugar composition during maturation of zygotic embryos

were observed with sucrose, galactinol, rafﬁ nose, verbascose, and

stachyose being the most prominent in mature seeds. In contrast,

pea somatic embryos contained much lower total soluble sugars

being composed of fructose, glucose, myo-inositol, sucrose, rafﬁ -

nose, and galactinol, but lacking stachyose and verbascose. Most

interestingly, irregular misshaped somatic embryos differed in their

carbohydrate proﬁ les from normal ones [

48 ]. Taken together,

these analyses on carbohydrates point to the fact that somatic

embryos often contained lower total amounts of soluble sugars, in

later stages show different monosaccharide:sucrose ratios and a

lack or smaller amounts of rafﬁ nose and its derivatives that are con-

sidered to be important for desiccation tolerance. Thus, matura-

tion obviously is the major bottleneck for somatic embryogenesis

in several species.

Traud Winkelmann

35

Comparative lipid analyses in both types of embryos are hardly

found in literature, except one report for Prunus avium [

49 ]: the

lipid proﬁ les of somatic embryo s resemble those of zygotic embryo s

with neutral glycerolipids and phosphatidylcholine being the major

lipid classes. However, contents of these two classes of lipids in

somatic embryos were comparable to those of immature zygotic

embryos, which was in line with the observation that somatic

embryos did not develop further, until they received a cold treat-

ment that resulted in increased lipid levels.

Polyamines (among which the commonly occurring spermidine,

spermine, and putrescine) are assumed to play a role in embryo-

genesis [

50 ] and they were quantiﬁ ed in somatic and zygotic

embryo s of Norway spruce [

51 ]. If mature somatic embryo s are

contrasted to zygotic ones, the latter contained less spermidine,

but more putrescine resulting in a much lower spermidine:putrescine

ratio. This ratio as well as the higher absolute polyamine contents

of somatic embryos may be connected to the lower germination

ability of somatic embryos. However this assumption requires

physiological explanations [

51 ] .

In a comparison of plant hormone contents in somatic and zygotic

larch embryos, 100 times higher concentrations of ABA were

found in somatic embryo s that were cultivated on medium con-

taining the nonphysiological ABA concentration of 60 μM. During

maturation the ABA content increased in somatic embryos while it

declined in zygotic ones [

52 ]. Among the cytokinins, only for iso-

pentenyladenine differences were detected with much higher levels

in zygotic embryo s, whereas IAA contents were similar in both

embryo types [

52 ]. The set of enzymes detoxifying reactive oxy-

gen species differed between zygotic and somatic embryos of horse

chestnut [

53 ]: catalases and superoxide dismutases showed differ-

ent courses of expression and different isoforms, especially in the

maturation phase that resembled more the germination phase in

case of somatic embryos. These authors concluded that somatic

embryos seem to be exposed to higher stress levels than their

zygotic counterparts.

While an increasing number of studies on gene expression during

embryogenesis of either the somatic (e.g. soybean, [

54 ]) or the

zygotic type (e.g. loblolly pine, [

55 ]), are available, only very few

reports deal with transcriptomic comparisons of somatic and

zygotic embryo s. In C. persicum, Hoenemann et al. [

27 ] com-

pared zygotic and somatic embryo s and also embryogenic and

nonembryogenic cell lines using a cDNA microarray with 1216

transcripts. They observed an upregulation of oxidative stress

response genes in somatic embryos, as for glutathione S-transferases,

catalase, and superoxide dismutase. These genes were upregulated

not only in early stages of somatic embryogenesis but also 3 weeks

2.2.3 Lipids

2.2.4 Polyamines

2.2.5 Plant Hormones

2.3 Comparison

of Transcriptomes

Somatic Versus Zygotic Embryogenesis

36

after induction, pointing at lingered stress and/or the induction of

secondary somatic embryos. The importance of pectin-mediated

cell adhesion as a prerequisite for embryogenicity was proposed by

these authors based on the higher abundance of several genes

encoding pectin-modifying enzymes in embryogenic than in non-

embryogenic cells. Moreover, a cationic peroxidase that prevents

cell expansion was suggested to be important for early embryogen-

esis [

27 ]. Thus, the early cell divisions that do not result in expan-

sion in size in early zygotic embryogenesis could be realized in a

similar way in somatic embryos.

Recently, next generation sequencing was applied in cotton to

compare the transcriptome of three comparable stages of both

somatic and zygotic embryo s [

56 ]. Among a total of more than

20,000 unigenes, 4242 were found to be differentially expressed in

these six samples. Of the differentially expressed genes a higher

number was upregulated in somatic embryo s at all stages [

56 ].

Especially, stress response genes including hormone-related genes

(mainly ABA and jasmonic acid signaling), kinase genes, transcrip-

tion factor s, and downstream stress responsive genes—e.g. late

embryogenesis abundant (LEA) genes, heat shock proteins—were

found at higher expression levels in somatic embryos. Moreover,

cotton somatic embryos were found to be metabolically more

active than their zygotic counterparts as indicated by gene expres-

sion data, the number of mitochondria , bigger vacuoles, and more

lipid droplets [

56 ]. Stress on the one hand can be considered as an

important trigger of embryo development which also occurs in the

zygotic system during maturation to prepare the embryo for desic-

cation stress. On the other hand, if cells experience too much stress

as it might be the case under in vitro conditions, this might disturb

the developmental program or even lead to cell death.

The proteome reﬂ ects the total set of proteins that is present in a

deﬁ ned tissue in a speciﬁ c developmental stage under deﬁ ned con-

ditions and thus provides direct evidence of the biochemical and

physiological status of these cells. A possible disadvantage of pro-

teomic studies is that proteins of very low abundance such as

important transcription factor s may be not detected. Although the

number of proteins that can be detected is limited if gel-based pro-

teomics is used, the comparison of two proteomes can be visualized

very well using 2D-SDS-PAGE (two-dimensional isoelectric focus-

sing/sodium dodecylsulfate polyacrylamide gel electrophoresis ).

In our own comparative studies we used a gel-based proteomic

comparison of somatic and zygotic embryo s of C. persicum , the

work-ﬂ ow of which is depicted in Fig.

2 [ 57 ]. The ﬁ rst and essen-

tial step is to select the biological material that will allow a mean-

ingful proteomic comparison; in our studies, the selection of

comparable stages was based on embryo morphology [

42 , 58 , 59 ].

Spots of interest being either more abundant or even speciﬁ c for

2.4 Comparison

of Proteomes

Traud Winkelmann

37

Fig. 2 Workﬂ ow of a gel-based proteomic approach combined with mass spectrometry . The biological system

represents one or more samples to be analyzed via a gel based proteomic approach. In the example given in

this diagram, proteomes of zygotic and somatic embryo s of Cyclamen persicum are analyzed and compared

( a ). Therefore, total proteins are extracted from each tissue ( b ) and separated via IEF-SDS PAGE ( c ). To perform

statistical analyses with gels of different tissues, at least a set of three replicates for each tissue is required.

Spots that differ signiﬁ cantly in abundance are labeled ( green and red ) in an overlay image of all gels analyzed

( d ). Protein of interest (e.g., differentially abundant proteins) are isolated from 2D gels and subsequently a

tryptic protein digest is performed ( e ). The resulting peptides are separated via liquid chromatography (LC)

before tandem mass spectrometry analyses ( f ). Protein identiﬁ cation is performed based on resulting peptide

sequences ( pink ) via a database search matching to known sequences ( g ). Finally, a digital proteome reference

map can be designed indicating all identiﬁ ed proteins ( h ). Using a gel-free shotgun approach, the steps ( c ) and

( d ) are replaced by digestion of a complex protein sample which is then further analyzed (reproduced from

[ 57 ] with permission from author and Leibniz Universität Hannover)

Somatic Versus Zygotic Embryogenesis

38

one sample can then be eluted from the gel and subjected to mass

spectrometry in order to identify the protein or proteins within

this spot by comparison to databases. Finally, the obtained data can

be combined to an interactive reference map which in our case was

made publicly accessible and allows ﬁ ltering spots by their abun-

dance, metabolic function, or tissue speciﬁ city [

60 ]. This tech-

nique was already applied in the ‘90s. Comparing somatic and

zygotic embryos of D. carota torpedo shaped somatic embryo s had

a clearly distinct protein pattern from zygotic embryos and lacked

the maturation speciﬁ c proteins, namely two globulin-type storage

proteins and a LEA protein [

61 ]. In the gymnosperm species

Norway spruce ( Picea abies ), similar protein patterns of zygotic

and somatic embryos, the latter cultivated on maturation medium

containing 90 mM sucrose and 7.6 μM ABA , were reported and

both types were dominated by storage proteins [

62 ].

Our model to study somatic embryo genesis is the ornamental

plant C. persicum . In a pilot study, the proteomes of cyclamen

somatic embryos grown in differentiation medium with 30 and 60

g/L sucrose were compared to zygotic embryo s and endosperm

[

42 ]. When somatic embryos were differentiated in medium con-

taining 60 g/L sucrose, 74 % of the protein spots were found in

comparable abundance as in the zygotic embryos’ proteome , while

11 % and 15 % were found in higher abundance in zygotic and

somatic embryos, respectively. Enzymes of the carbohydrate

metabolism, as well as heat shock proteins and a glutathione-S-

transferase, were more abundant in somatic embryos. Thus, again

evidence was presented for differences in stress response of both

types of embryos. Furthermore, ﬁ rst insights into cyclamen seed

storage protein accumulation and the synthesis of the storage car-

bohydrates xyloglucans were gained [

42 ] . A follow-up study made

use of the advances achieved in protein extraction, resolution, eval-

uation, more sensitive mass spectrometrical analyses and, most

important, sequence information available in the data bases leading

to higher identiﬁ cation rates even for this nonmodel organism

[

58 ]. In both embryo types glycolytic enzymes were identiﬁ ed as a

high percentage of the identiﬁ ed proteins. In somatic embryos

four protein spots showed six- to more than tenfold increased

abundance, and the identiﬁ ed proteins within these spots were

involved in oxidative stress defense: osmotin -like protein and anti-

oxidant 1, peroxiredoxin type 2, and catalase. This ﬁ nding is a clear

indication that somatic embryos are much more stressed than

zygotic ones [

58 ]. The occurrence of truncated forms of enolases

in zygotic embryos in relatively high amounts that disappear dur-

ing germination suggested a new role of parts of this glycolytic

enzyme as storage proteins [

58 ] . We followed the original idea of

taking the proteome of zygotic embryos as a reference for the opti-

mized development of high quality somatic embryos: we could

show that in somatic embryos a change of the proteome towards

Traud Winkelmann

39

the zygotic status was induced after the application of a maturation

treatment with ABA [

59 ]. After ABA treatment, the proposed new

storage proteins (“small” enolases) appeared in the proteome of

somatic embryos, thus resembling more the proteome of zygotic

embryos (Fig.

3 ). Sghaier-Hammami et al. [ 64 ] found the total

Fig. 3 Upper Part: Comparison of protein gels of torpedo-shaped somatic embryo s, zygotic embryo s, and

somatic embryos treated with 10 mg/L ABA for 28 days (taken from different studies [ 57 , 63 ], encircled are

parts of the gels which show high similarity in zygotic and ABA-treated embryos). Lower Part: Alterations in

protein abundance of 56 days old somatic embryos after cultivation on medium containing 0, 2, and 10 mg/L

ABA for 28 days. Green labeled spots are at least 1.5 times higher abundant in controls, orange labeled spots

are at least 1.5 times more abundant in the 2 mg/L ABA treatment, and pink labeled spots are at least 1.5

times more abundant in the 10 mg/L ABA treatment (compared to control) (lower part of the ﬁ gure reproduced

from [ 63 ] with permission from the author and Leibniz Universität Hannover)

Somatic Versus Zygotic Embryogenesis

40

protein content as well as the number of spots to be higher in

zygotic than in somatic embryos of date palm in a comparative

2-DE proteomic approach. Sixty percent of the protein spots dif-

fered in their abundance between the two embryo types, and out

of 63 spots of differential abundance that were eluted from the

gels, 23 were identiﬁ ed. Most of the proteins of higher abundance

in somatic embryos were involved in the glycolysis pathway, citrate

cycle, and ATP synthesis pointing to a higher energy demand,

while in zygotic embryos a high abundance of storage proteins and

stress-related protein s of the heat shock family indicated matura-

tion and preparation of dehydration [

64 ].

Also in cocoa, enzymes of the carbohydrate and energy metab-

olism were very prominent in torpedo stage somatic and zygotic

embryo s [

65 ]. Interestingly, somatic embryo s had a more active

oxidative/respiration pathway while in zygotic embryos anaerobic

fermentation might be the more important energy pathway. Again

stress-induced proteins such as peroxidases , pathogenesis-related

proteins, and glutathione S-transferase were more abundant in

somatic embryos [

65 ].

3 Role of the Endosperm

Somatic embryo s lack an endosperm , which is not only a tissue that

nourishes the developing embryo and the germinating seedling,

but insulates the embryo from mechanical pressure and has impor-

tant signaling function for embryo development , maturation and

growth arrest, and ﬁ nally germination timing [

66 ]. Thus, for opti-

mization of somatic embryo genesis a detailed look into the endo-

sperm during seed development seems reasonable.

In order to develop optimal culture media for somatic embryo

development in wheat , Carman et al. [

67 ] analyzed minerals and

primary metabolites of the endosperm during seed development.

Maltose concentrations in the extracted kernel ﬂ uid increased

between 6 and 18 days after pollination indicating that this product

of starch hydrolysis is the major carbon source for the developing

embryo. For the development of improved tissue culture media,

the addition of free amino acids, the adjustment of phosphate and

sulfur which were detected in relatively high concentrations in the

kernel ﬂ uids probably because of their presence in phosphorylated

sugars and amino acids, respectively, and the addition of maltose

and short chain fructans were suggested [

67 ]. Likewise in white

spruce , somatic and zygotic embryo s and the megagametophyte

which is the haploid nourishing tissue of gymnosperms were ana-

lyzed with respect to their mineral contents [

68 ] . The female game-

tophytes and zygotic embryo s contained more phosphorus,

potassium, magnesium, and zinc on a dry- weight basis than somatic

embryo s, whereas the female megagametophyte stood out due to

Traud Winkelmann

41

its high calcium content when compared to the embryo tissues

[

68 ]. However, if this information is going to be integrated into

optimization of culture media, more data sets will be necessary for

the mineral contents in different developmental phases, and also

the forms in which the minerals are found in the respective tissue.

Arabinogalactan protein s were identiﬁ ed in conditioned culture

media of embryogenic cells by Kreuger and van Holst [

69 ] and

found to be essential for somatic embryo development [

70 ]. Most

interestingly, an endochitinase gene (EP3) which is involved in the

generation of arabinogalactan protein s was expressed in carrot

seeds by cells in the integuments and the protein localized in the

endosperm and also in nonembryogenic cells of embryogenic cul-

tures [

70 ]. Also the formation of arabinogalactan proteins in the

developing carrot seed was shown to be developmentally regulated

[

71 ]. In a review Matthys-Rochon [ 72 ] came to the conclusion

that nonembryogenic cells within embryogenic cultures might

take over some functions of the endosperm by secretion of signal

molecules that control embryo development.

For C. persicum the proteomic analysis of the endosperm dur-

ing seed development revealed a general shift from high molecular

weight proteins to low molecular weight proteins and the accumu-

lation of storage proteins (including “small” enolases) from 7

weeks after pollination when the endosperm is still liquid [

73 ].

Furthermore proteins involved in synthesis of other storage com-

pounds, namely lipids and xyloglucans were identiﬁ ed in the endo-

sperm. Obviously, stress response including reactive oxygen species

detoxiﬁ cation and ABA signaling also play a role in endosperm and

embryo development [

73 ].

4 Conclusions and Outlook

It can be concluded from the aforementioned literature that:

1. Somatic embryo s are more exposed to stress than their zygotic

counterparts,

2. Somatic embryo s accumulate less storage compounds,

3. Somatic embryo s do not undergo a proper maturation phase

that would include a growth arrest but instead germinate

precociously.

The role of stress which is on the one hand an important trig-

ger of embryogenesis and, on the other hand, induces severe

changes in the cellular metabolism; here especially the role of reac-

tive oxygen species deserves further investigations. Obviously, par-

ticularly somatic embryo genesis is a process that only is successfully

realized if the cells experience the right stress level at the right

developmental time frame. Also programmed cell death which has

Somatic Versus Zygotic Embryogenesis

42

an impact in zygotic and somatic embryogenesis should be taken

into consideration in coming research projects. The importance of

the maturation phase for accumulation of storage reserves , and

also for the clear distinction of differentiation and germination , has

been noticed in many systems. Nevertheless, input is needed par-

ticularly to improve this phase of somatic embryogenesis in the

future. At physical culture conditions, attention is not often paid,

except at the oxygen concentration, for example in wheat embryo-

genesis [

3 , 74 ]. Here it has been shown that installing reduced O

2

levels, mimicking the situation found in seeds, improved growth

and development of somatic embryos. However, the O

2 levels

changed not only with time of development and spatially but also

during the day due to photosynthesis [

74 ]. Our own studies in

cyclamen revealed hypoxic conditions in seeds at the position

where the embryo is found about 5–6 weeks after pollination in

unpublished measurements according to [

75 ]. Thus, in vitro cul-

tured somatic embryos which grow at ambient oxygen concentra-

tions may establish too high or altered metabolic activity as

indicated by some studies cited above (e.g. [

64 ] , [ 65 ]) and/or

oxidation of plant growth regulator s such as cytokinins, ABA , and

indole acetic acid due to increased activity of oxidases as discussed

by Carman and Bishop (2004) [

74 ] .

The “omics” tools (transcriptomics, proteomics, metabolo-

mics…) will substantially improve in terms of sensitivity, resolution

and identiﬁ cation, and affordable analyses of different genotypes

over time and thereby enable us to gain deeper insights into plant

embryogenesis and to optimize the in vitro protocols for somatic

embryo genesis. Moreover, epigenetic regulation of embryogenesis

by methylation/demethylation and histone modiﬁ cations , post-

transcriptional and posttranslational modiﬁ cations should be stud-

ied in detail especially during the early phases. The role of speciﬁ c

micro RNAs as regulators of plant development including embryo-

genesis has to be elucidated, since Oh et al. [

76 ] found differences

in the abundance of ﬁ ve micro RNAs between somatic and zygotic

embryo s in loblolly pine. Although zygotic embryogenesis is more

and more understood because of mutant analyses and molecular

genetic studies of embryogenesis-related genes and both kinds of

embryogenesis are studied in detail on a transcriptional and pro-

teomic level, many aspects of the fascinating regeneration pathway

of plant embryogenesis are still not explained. One interesting

aspect for instance is the fact that somatic embryogenesis is highly

dependent on the genotype, whereas zygotic embryogenesis is not.

Especially for the recalcitrant genotypes improvements would be

desirable by learning from seeds.

Traud Winkelmann

43

Acknowledgements

Special thanks go to Dr. Dolf Weijers and Dr. Jos Wendrich and

John Wiley and Sons (publisher) for the permission to use Fig.

1 in

this book chapter. The author is very grateful to Dr. Christina

Rode and MSc. Kathrin Lindhorst for their allowance to publish

Figs.

2 and 3 and for the very good scientiﬁ c cooperation. The

excellent proofreading of the manuscript by Dr. Melanie Bartsch

and Dr. Bernhard Strolka is gratefully acknowledged.

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_3, © Springer Science+Business Media New York 2016|

Chapter 3

Recent Advances on Genetic and Physiological Bases

of In Vitro Somatic Embryo Formation

Maria Maddalena Altamura , Federica Della Rovere , Laura Fattorini ,

Simone D’Angeli , and Giuseppina Falasca

Abstract

Somatic embryogenesis involves a broad repertoire of genes, and complex expression patterns controlled

by a concerted gene regulatory network. The present work describes this regulatory network focusing on

the main aspects involved, with the aim of providing a deeper insight into understanding the total repro-

gramming of cells into a new organism through a somatic way. To the aim, the chromatin remodeling

necessary to totipotent stem cell establishment is described, as the activity of numerous transcription fac-

tors necessary to cellular totipotency reprogramming. The eliciting effects of various plant growth regula-

tors on the induction of somatic embryogenesis is also described and put in relation with the activity of

speciﬁ c transcription factors. The role of programmed cell death in the process, and the related function of

speciﬁ c hemoglobins as anti-stress and anti-death compounds is also described. The tools for biotechnol-

ogy coming from this information is highlighted in the concluding remarks.

Key words Auxins , Chromatin remodeling , Hemoglobins , Osmotin , Programmed cell death ,

Receptor-kinase s , Stem cells , Stress signaling , Somatic embryo s , Transcription factor s

1 The Concept of Stem Cell in Somatic Embryogenesis

1

Most plant cells express developmental plasticity allowing their

reprogramming. The developmental plasticity is linked to the stem

cell condition, because the stem cells have the unique characteristics

1 Genetics nomenclature adopted in the chapter:

– Wild-type gene names are uppercase italic, e.g., EMBRYOMAKER.

– Wild-type gene symbols consist of three uppercase letters in italics, e.g., EMK , and may be preceded by

two letters in italics, the ﬁ rst one in uppercase, showing the species to which the gene belongs, e.g.,

AtEMK means Arabidopsis thaliana EMBRYOMAKER .

– Mutant gene names are lowercase italic, e.g., embryomaker .

– Mutant gene symbols are designed by three lowercase letters in italics corresponding to the gene locus

name, e.g., emk .

– Different genes with the same symbol are distinguished by different numbers, e.g., LEC1 and LEC2 .

– Different alleles of the same gene are distinguished with a number following a hyphen, e.g., Hb1-2 .

48

to both self-renew and develop into precursors that can form

different cell types and tissues. During zygotic embryo formation

the number of stem cells reduces progressively, and the stem cells

become limited to the opposite poles of the mature embryo,

forming niches in the shoot and root meristems. The zygote, and

its early derivatives, are totipotent stem cells, whereas the niches

in the root and shoot meristems of the mature embryo are formed

by pluripotent stem cells. Pluripotent stem cells are also present

out of the apical meristems, e.g. in the procambium, cribro-vas-

cular cambium and phellogen, and in the meristemoids, as those

originating stomata and hairs ([

1 ] and references therein). Plant

somatic embryo genesis (SE) is the process through which differ-

entiated cells, single or in small groups, reverse their develop-

mental program during in vitro culture, and rarely in planta ,

giving rise to embryos which follow a developmental pattern

similar to the one of zygotic embryogenesis [

2 ]. The cells that

initiate the somatic embryo, and their early derivatives (trans-

amplifying cells), have been recently included in the plant stem

cell concept [

3 ]. Moreover, stem cell niches are maintained in the

apical meristems of the somatic-embryo-derived plants, and are

pluripotent as in zygotic embryo-derived plants [

1 ]. However,

the mechanisms underlying the initiation of somatic embryos are

still poorly understood [

4 , 5 ]. Deciphering the molecular deter-

minants of SE can contribute to revealing the genetic program

underlying the phenomenon of stem cell totipotency and pluri-

potency, and somatic embryo formation and maturation. The

study of regulatory molecules and associated gene networks dur-

ing SE is essential for understanding embryogenic competence

and plant regeneration, which are necessary for crop improve-

ment and the establishment of new protocols, e.g., aimed to the

production of synthetic seeds and the maintenance of elite

germoplasm.

2 Chromatin Remodeling as a Prerequisite for Totipotent Stem Cell

Establishment in SE

In animals, chromatin remodeling is an important tool of stem cell

conversions [

6 ]. In plants, chromatin structure is continuously

remodeled during development, whereas a chromatin-dependent

gene silencing is a common mechanism for maintaining the

– Proteins are written in non-italic uppercase, with the same full descriptive name, and the symbol

of the corresponding gene, e.g., EMBRYOMAKER, and EMK.

– The symbol of either a gene or a protein known by multiple names is given by the known symbols sepa-

rated by a slash, e.g., BAK1/SERK3 and BAK1/SERK3, respectively.

Maria Maddalena Altamura et al.

49

differentiated cellular state. Thus, as in animals, a role of chromatin-

structure changes in pluripotency of plant stem cells has been

proposed [

7 – 9 ]. The stem cells show a more dynamic chromatin

state than the differentiated cells because structural chromatin pro-

teins, like histones, are exchanged more rapidly in stem cells than

in differentiated cells, and this favors rapid changes in the gene

expression program [

10 ].

It has been hypothesized that chromatin restructuring plays

two major roles during early stages of SE. An unfolding of the

super-coiled chromatin structure is required for the dedifferentia-

tion of the somatic cells, both when they produce embryos directly

(direct SE), and when they are engaged into callus formation

before embryogenesis (indirect SE). This early step is necessary to

allow the expression of genes which had been inactivated by het-

erochromatization during the cell differentiation process, but

which are speciﬁ cally required for the embryogenic pathway [

7 ]. A

chromatin remodeling is also required at the end of embryogenic

process to repress the embryo-speciﬁ c genes, thus reactivating the

differentiation process, at least in speciﬁ c cellular districts of the

somatic embryo s. Formation of somatic embryos by somatic

embryo cells (i.e., secondary SE) occurs when the repression of the

embryo-speciﬁ c gene pathway and the chromatin re-folding are

delayed/altered, not allowing the transition from the embryo to

the seedling. In plants, as well as in animals, two major epigenetic

pathways play important roles in the regulation of cell-fate deci-

sions by modifying chromatin, namely DNA methylation and his-

tone methylation. In animals, there is increasing evidence for a

complex regulatory interplay between the two pathways, with a

direct mechanistic link based on physical interactions of histone-

modifying proteins and DNA methyltransferases ([

11 ], and refer-

ences therein).

Polycomb group (PcG) proteins are present in animals and

plants. They maintain the inactive state of the target genes by

establishing repressive histone modiﬁ cations , e.g. by methylation.

During Arabidopsis life cycle, distinct variants of the POLYCOMB

REPRESSIVE COMPLEX 2 (PRC2) are involved in the regula-

tion of the developmental processes, such as gametophyte and seed

development and embryo-to-seedling transition [

11 ]. The plant

variants of PRC2 catalyze the Histone H3 Lysine 27 Trimethylation

(H3K27me3) [

12 ]. It is known that H3K27me3 is a repressive

mark that plays a crucial role in the dynamic regulation of gene

expression in plant development [

13 ], acting as a major silencing

mechanism. In Arabidopsis , prc2 mutants with substantially

reduced levels of H3K27me3 exhibit extensive derepression of the

embryonic traits ([

14 ], and references therein). The ATP-

dependent chromatin remodeler PICKLE (PKL) belongs to a pro-

tein family that can participate in multiple remodeling pathways

Genetics and Physiology of Somatic Embryogenesis

50

and can either repress or activate gene expression depending on

the other factors it associates with. In Arabidopsis , PKL promotes

the epigenetic mark H3K27me3 facilitating repression of speciﬁ c

genes. In fact, the phenotype of pickle ( pkl ) mutant is characterized

by the postembryonic expression of embryo-speciﬁ c markers, e.g.

the LEAFY COTYLEDON ( LEC ) genes (see below), and by the

spontaneous regeneration of somatic embryo s in the roots [

15 ].

Thus, the loss of PKL generates a window of opportunity through-

out which the embryo transcriptional program has the potential to

become reestablished [

14 ], maintaining the embryo-related gene

expression.

In contrast with the effects of histone methylation, DNA

(hyper)methylation, for example caused by exogenous auxin,

which is, in general, necessary to induce SE, is positively related to

SE ([

16 ], and references therein) (Fig. 1 ).

Fig. 1 Model of chromatin remodeling involved in SE-induction. 2,4-D-activated DNA hypermethylation , his-

tone (H) demethylation and hyperacetylation events, and early-activated/repressed proteins are shown (see

the text for further explanation). Numbers (1) and (2) consequences of PKL knock-out

Maria Maddalena Altamura et al.

51

In Arabidopsis , DNA methylation is mediated by at least three

classes of methyltransferases. The METHYLTRANSFERASE1

(MET1) is one of these classes. Zygotic embryo s with loss-of-

function mutations in MET1 develop improperly, displaying altered

cell divisions, reduced viability, mis-expression of genes specifying

embryo cell identity, and altered auxin hormone gradients [

17 ]. In

carrot , a MET1 gene is expressed transiently after the application of

the synthetic auxin 2,4- D , and before the formation of the SE-cell-

clumps, and 5-azacytidine, an inhibitor of DNA methylation, sup-

presses the embryogenic clump formation [

18 ]. All together, these

results highlight a role of MET1 in both zygotic and somatic

embryo genesis. Very recently, it has been observed that during

seed development Arabidopsis MET1 interacts with MEDEA, one

of the core components of the FERTILIZATION INDEPENDENT

SEED (FIS)-PRC2 complex, with MEDEA involved in PRC2

repression ([

12 ], Fig. 1 ). The interaction between MET1 and

MEDEA (Fig.

1 ) demonstrates, for the ﬁ rst time in plants, that a

concerted action of the epigenetic pathways of DNA methylation

and histone methylation regulates the switching of developmental

changes [

11 ], and sustains that a concerted action of DNA meth-

ylation and histone demethylation might be essential for SE induc-

tion (Fig.

1 ). The modiﬁ cations of chromatin also include the

acetylation of histone tails for relaxing the packing of the DNA,

thus facilitating the access to DNA of many regulatory proteins.

Thus, the hyperacetylation of histones is associated with active

gene expression, while hypoacetylation correlates with gene repres-

sion ([

19 ], and references therein). In animals, a dynamic repro-

gramming of both histone acetylation and methylation has been

demonstrated, e.g. in cloned mouse embryos [

20 ], whereas full

evidence of this interplay is still lacking for plant zygotic/somatic

embryogenesis.

Proteins containing bromodomains have the role of decipher-

ing the histone acetylation codes. There are bromodomain pro-

teins that also contain an Extra Terminal domain that is a

protein-protein interaction motif [

21 ]. BROMODOMAIN and

EXTRA TERMINAL DOMAIN proteins (BET proteins) bind to

acetylated lysines of histone tails and control gene transcription

([

22 ], and references therein) (Fig. 1 ). In a variety of organisms

the BET proteins contribute to the transmission of the transcrip-

tional memory from one generation of cells to the next ([

23 ], and

references therein). In Arabidopsis , the BET bromodomain factor

GENERAL TRANSCRIPTION FACTOR GROUP E4 (GTE4)

is involved in the activation and maintenance of cell division in the

meristems. The loss of GTE4 negatively affects both zygotic

embryo genesis by altering meristem organization in the mature

embryo, e.g., at the root pole (Fig.

2a, b ), and post-embryonic

growth by altering the stem cell niche formation in the root apical

meristems [

22 , 23 ]. The RETINOBLASTOMA (Rb)-E2

Genetics and Physiology of Somatic Embryogenesis

52

Fig. 2 Arabidopsis zygotic embryo s ( a – b ). Altered root pole meristem in a mature gte4 embryo ( a ), in compari-

son with the well-organized meristem of the wild-type ( b ). ( c – d ) Auxin transport and localization in the mature

embryo. The expression pattern of PIN1 auxin-efﬂ ux carrier in a PIN1::GUS embryo [i.e., a transgenic embryo

expressing the uidA gene coding for a β - GLUCURONIDASE (GUS) under the control of PIN1 promoter] ( c ), and

Maria Maddalena Altamura et al.

53

FACTOR (E2F) pathway is considered an essential link between

chromatin restructuring, dedifferentiation, and fate switch. In

tobacco protoplasts, the Rb/E2F-target genes RNA RIBOSOMAL

CLUSTER 2 ( RNR2 ), i.e., the small subunit of ribonucleotide

reductase, and PROLIFERATING CELL NUCLEAR ANTIGEN

( PCNA ) are condensed and silent in differentiated leaf cells, but

become de-condensed as cells acquire competence for fate-switch,

and turn transcriptionally active during progression into S phase,

concomitantly with Rb phosphorylation [

24 ]. Moreover, Rb has

been shown to bind to a transcription factor (TF) that functions in

the Arabidopsis root stem cell niche, and a relationship with

WUSCHEL (WUS)/CLAVATA (CLV) has been also shown

([

19 ], and references therein). Roles for Rb proteins in human

embryogenesis are widely known [

25 ]. GTE4 might be a good

candidate in the control of histone-acetylation during plant SE,

because, as the Bromodomain containing 2 (BRD2/RING3) pro-

tein in animals, its activity is related to the Rb-E2F pathway [

22 ]

(Fig.

1 ). Among the possible factors causing chromatin modiﬁ ca-

tions there are also small RNAs, such as the small-interfering RNAs

(siRNAs) and the microRNAs (miRNAs). The small RNAs not

only function at the posttranscriptional level by guiding sequence-

speciﬁ c transcript degradation and/or translational repression

([

26 ], and references therein), but can also play a role in targeting

DNA methylation through RNA-directed DNA methylation [

27 ,

28 ]. These events lead to chromatin modiﬁ cations eventually

resulting in transcriptional silencing and heterochromatin forma-

tion [

29 ]. ARGONAUTE (AGO) effectors of RNA silencing bind

small RNA molecules and mediate mRNA cleavage, translational

repression, or DNA methylation [

30 ]. The possible regulation of

SE by small RNAs has been investigated using various systems. In

rice callus, a unique set of miRNAs, only expressed or differentially

expressed in embryogenic cells, was identiﬁ ed [

31 , 32 ]. Moreover,

miRNA expression during SE was also characterized in orange

[

33 ], longan [ 34 ], and cotton, where four trans-acting small inter-

fering siRNAs (tas3-siRNAs) were also identiﬁ ed [

35 ]. On the

other hand, the expression of AGO gene s during SE is well known,

and in both gymnosperms and angiosperms , e.g., spruce [

36 ], car-

rot [

37 ], Cichorium intybus [ 38 ], Auracaria angustifolia [ 39 ].

Nonetheless, the roles of miRNAs and siRNAs, and related effec-

tors, in the induction of plant SE remain to be understood.

Fig. 2 (continued) the strictly apical localization of auxin in the root pole of a DR5::GUS embryo [i.e., a trans-

genic embryo expressing the GUS -encoding reporter gene under the control of the synthetic auxin-responsive

promoter DR5 ] ( d ), are shown. ( e – f ) Calcium distribution in the embryo monitored by the CTC-Ca

2+ epiﬂ uores-

cence signal ( yellow - green color ). The apical-basal distribution along the hypocotyl ( e , arrow ), and the strong

signal-reduction at the root apex ( f ), are shown. Bars = 10 μm ( a , b , d – f ) and 50 μm ( c )

Genetics and Physiology of Somatic Embryogenesis

54

3 Transcription Factor Activity Involved in Cellular Totipotency Reprogramming

The developmental cell switching to SE induction involves activa-

tion of various signal cascades and differential gene expression.

The inductive role of Plant Growth Regulators (PGRs), auxin in

particular, has been well established, and will be summarized

below. However there is increasing evidence for a role of numerous

Transcription Factors (TFs) in accordance with human somatic

cells in which a speciﬁ c combination of TFs re-programs the dif-

ferentiated cells into embryonic stem cells [

40 ]. In plants, TFs

involved in SE induction have been reported. For example, an

extensive modulation of the TF- transcriptome has been recently

described during SE induction by in vitro culture in Arabidopsis sug-

gesting directions for further research on functional genomics of SE

[

41 ]. In this model plant, it has been demonstrated that the embryo-

induction stage is associated with a robust change of the

TF-transcriptome by a drastic upregulation of transcripts related

with plant development, PGRs and stress response s. By contrast, the

advanced embryo stages are associated with the stabilization of the

transcript levels of the majority of the TFs [

41 ]. TFs are known to

play fundamental roles in the control of plant cell totipotency , which

is essential for SE (see above), and the list of TFs affecting SE induc-

tion includes BABY BOOM ( BBM ) [

42 ], WUS [ 43 ], some

WUSCHEL RELATED HOMEOBOX genes ( WOX ) [

44 ],

AGAMOUS-LIKE15 ( AGL15 ) [

45 ], LEAFY COTYLEDON ( LEC )

[

46 ], genes encoding MYELOBLASTOSIS (MYB) TFs, i.e.,

AtMYB115 and AtMYB118 [

47 ], and EMBRYOMAKER [ 48 ].

BBM was isolated from microspore embryo cultures of Brassica

napus [

42 ]. It belongs to a family of genes [ APETALA2

( AP2 )/ ETHYLENE RESPONSE FACTOR ( ERF )] known to

enhance regeneration in vitro, and to be involved into meristem

cell fate and organ development. Interestingly, in Arabidopsis ,

BBM induces somatic embryo formation from seedlings in the

absence of exogenously applied PGRs ([

49 ], and references

therein). Arabidopsis EMBRYOMAKER ( AtEMK ) is another

AP2-domain TF. It is homologous to BnGemb-18 of Brassica

napus which is speciﬁ cally expressed in microspore embryogenesis ,

as BBM ([

48 ], and references therein). AtEMK ectopic expression

results into embryo-like structures from cotyledons in planta ,

enhancing somatic embryogenesis in in vitro culture, and a role for

this gene in conferring embryonic identity to cells has been pro-

posed [

48 ].

Another initiator of ectopic embryogenesis is AGL15 . This

gene was identiﬁ ed from Arabidopsis and soybean as a MADS

domain-containing TF speciﬁ cally expressed in the embryonic cells

[

50 , 51 ]. The MADS name derives from the initials of the found-

ing members of the gene family, i.e., MINICHROMOSOME

Maria Maddalena Altamura et al.

55

MAINTENANCE1 ( MCM1 ), AGAMOUS ( AG ), DEFICIENS

( DEF ), and SERUM RESPONSE FACTOR ( SRF ). In plants, e.g.

Arabidopsis and rice, MADS-domain proteins are central players of

many developmental processes including ﬂ owering time, ﬂ oral

organogenesis, fruit and seed development ([

52 ], and references

therein). About embryogenesis, Harding and coworkers [

45 ] have

demonstrated that in Arabidopsis the ectopic expression of AGL15

enhances somatic embryo formation both from zygotic embryo s

removed from the seed at the green cotyledon stage and cultured

on a germination medium without exogenous PGRs, and from the

shoot apical meristem of seedlings growing in a liquid medium in

the presence of 2,4-D. It is also known that AGL15 enhances SE

reducing gibberellic acid (GA) levels by inducing a GA2-oxidase

which inactivates GA [

45 , 53 , 54 ]. Moreover, it is a component of

SOMATIC EMBRYOGENESIS RECEPTOR-LIKE KINASE 1

(SERK1) protein complex [

55 ], as detailed in a following

paragraph.

MYB proteins are TFs with a speciﬁ c DNA-binding domain

comprising up to three imperfect tandem repeats (R1, R2, R3),

that fold into a helix-turn-helix motif. In vertebrates, the MYB

gene family is small and includes c-MYB , A-MYB , and B-MYB ; the

products of these genes are involved in the control of cell prolifera-

tion, differentiation, and apoptosis [

56 ]. In plants, the MYB family

is much more extensive: at least 198 MYB genes have been identi-

ﬁ ed in Arabidopsis , and, similarly to the MADS -domain genes, are

involved in a wide range of developmental processes, including cell

cycle progression, cell differentiation, lateral organ polarity , ﬂ ower

and seed development. Plant MYB proteins are classiﬁ ed according

to the number of MYB repeats, and those with R2R3 repeats con-

stitute the largest group ([

57 ], and references therein). AtMYB118

and the closely related AtMYB115 encode R2R3-type MYB TFs.

In Arabidopsis their overexpression efﬁ ciently induces SE from

root explants, resulting in elevated expression levels of Arabidopsis

LEAFY COTYLEDON 1 ( LEC1 ) (see below), suggesting that

they may act as positive regulators of vegetative-to-embryonic

transition in a WUS -independent manner [

47 ], possibly acting

upstream to LEC1 .

The WUS gene , encoding a homeodomain protein, is critical

for stem cell fate determination in the shoot apical meristem of

higher plants. WUS activity results in signaling to the overlaying

stem cells , inducing CLAVATA3 ( CLV3 ). CLV3 acts as the ligand

for the CLAVATA1/CLAVATA2 (CLV1/CLV2) receptor com-

plex that limits the expression areas of WUS in the shoot apical

meristem, with this negative CLV3/WUS feedback loop ensuring

the shoot apical meristem homeostasis by regulating the number of

stem cells in the central zone [

58 , 59 ]. During zygotic embryo-

genesis in Arabidopsis WUS is expressed before the stem cell estab-

lishment in the embryonic shoot [

60 ]. In the same plant,

Genetics and Physiology of Somatic Embryogenesis

56

WUS -induced overexpression causes increased SE without any

external PGR , suggesting the involvement of this TF in the SE

process by promoting the vegetative-to-embryonic transition [

43 ].

Moreover, WUS induction occurs earlier than that of CLV3 , mark-

ing the initial cell clumps of SE. These clumps exhibit the same

cytological features of the pre-embryogenic aggregates formed in

Cyclamen persicum SE - callus, and which express other stem cell

markers which are not TFs (i.e., CpSERK1 and SERK2, as

described below) [

3 ]. The induction of WUS expression in the

embryogenic callus of Arabidopsis requires the removal of auxin

from the medium; however, the cell status in the embryonic callus

is regulated by auxin in a concentration-dependent manner, with

the levels of this exogenously applied PGR essential for determin-

ing WUS expression pattern. Moreover, the auxin gradients acti-

vate the polar auxin transporter PIN-FORMED 1 (PIN1) in the

embryogenic callus, and the suppression of both WUS and PIN1

show that both genes are necessary for somatic embryo induction

[

3 ] ( see the following paragraph).

The WUS-RELATED HOMEOBOX ( WOX ) gene family is a

class of TFs involved in the early phases of Arabidopsis zygotic

embryo genesis and in lateral and adventitious organogenesis [

44 ,

61 ]. In particular, WOX5 is a stem cell marker in the root apical

meristem and is very early expressed in in vitro adventitious root

induction ([

61 ], and references therein). Moreover, in Arabidopsis

WOX2, WOX8, and WOX9 are important cell fate regulators of

zygotic pro-embryos, acting as embryo-identity genes [

44 , 62 ].

During the SE-process WOX5 and WUS are expressed before the

embryo-identity genes, e.g. WOX2 [

63 ]. An extensive study on

the expression of WOX gene family in Vitis vinifera SE shows that

the WOX genes play important roles in coordinating the gene

transcription involved in the early phases of the process. VvWOX2

and VvWOX9 are the principal WOX genes expressed, and the low

aptitude to SE shown by speciﬁ c grape cultivars correlates with a

very low expression of these genes [

64 ].

Arabidopsis LEC gene s , LEC1 , LEC2 , and FUSCA3 ( FUS3 ),

were identiﬁ ed originally as loss-of-function mutations resulting in

defects in both embryo identity and seed maturation processes

[

65 ], and are all essential for SE induction in Arabidopsis [ 46 ].

LEC1 encodes a protein with sequence similarity to the HEME-

ACTIVATED PROTEINS 3 ( HAP3 ) subunit of the CCAAT bind-

ing factor [

66 ]. LEC1-LIKE ( LIL ) gene is also required for somatic

embryo development and is active in numerous plants. For exam-

ple, in cocoa, TcLIL mRNA levels are detected in young somatic

embryos and undetected in nonembryogenic explants and mature

somatic embryos, suggesting that also LEC-like genes may be

important in coordinating primary events leading to embryonic

competence ([

49 ], and references therein). LEC2- and FUS3-

proteins share greatest similarity with the B3 domain, a DNA-

Maria Maddalena Altamura et al.

57

binding motif unique to plant TFs, acting primarily in developing

seeds ([

46 ], and references therein). FUS3 activates transcription

of maturation-speciﬁ c genes containing RY domains [

67 ], and the

ectopic expression of LEC2 causes accumulation of lipids and seed

storage proteins in transgenic seedlings [

68 ]. In addition to its ver-

satile regulatory functions in zygotic embryo genesis and seed

development ([

69 ], and references therein), LEC2 , the same as

LEC1 , is sufﬁ cient to induce embryo development in vegetative

cells when expressed ectopically ([

49 , 68 ], and references therein).

Moreover, it directly induces genes involved in maturation pro-

cesses before formation of somatic embryos [

68 , 70 , 71 ]. The role

of LEC1 in maintaining embryonic characteristics in vegetative

organs requires auxin and sugar ([

72 ], and references therein), and

the capacity of SE in lec1lec2 double mutants is very low even in

the presence of auxin, suggesting that in Arabidopsis the formation

of somatic embryos by auxin needs the function of LEC genes

[

46 ]. By the use of an inducible chimeric fusion construct it has

been recently shown that ectopic expression of these LECs confers

embryonic characteristics also to tobacco [

72 ]. PKL, the chromatin-

remodeling factor described before, seems to be the master regula-

tor of LEC genes because, in the pkl mutant, roots express the

ability to form somatic embryos through a derepression of the

LEC genes [

73 ] (Fig. 1 ). Interestingly LEC2 overexpression leads

to spontaneous embryo formation in planta , but impairs SE

in vitro under auxin treatment, suggesting an auxin-mediated

mechanism of action [

70 ]. In line with this hypothesis, it has been

shown that LEC2 controls the INDOLE-3-ACETIC ACID

INDICIBLE30 ( IAA30 ), an auxin signaling gene, the PIN1 and

PIN2 auxin efﬂ ux carrier genes, the auxin biosynthesis YUCCA2

( YUC2 ) and YUCCA4 ( YUC4 ) genes, the AUXIN RESPONSE

FACTOR ( ARF ) genes ARF5 , ARF8 , and ARF10 , and SERK1 ,

the auxin-induced marker-gene of SE in a lot of plants ( see the next

paragraph) [

70 – 72 ]. Again in accordance, LEC2 is upregulated in

a SE culture induced on an auxin-containing medium, and the

gene overexpression compensates for the auxin treatment as

somatic embryos are formed in explants cultured under auxin-free

conditions [

74 ]. Very recently, it has been shown that de novo

auxin production via the tryptophan-dependent indole-3-pyruvate

(IPA)-YUC auxin biosynthesis pathway is implicated in SE induc-

tion, with LEC2 playing a key role in this mechanism [

69 ]. The

possibility that LEC2 may also promote SE due to a repression of

GA levels via a positive control on AGL15 , as well as of other

GA-related factors, has been also proposed [

69 , 71 , 72 ]. Moreover,

most genes involved in ethylene signaling pathway are downregu-

lated by LEC2 during SE in transgenic tobacco, suggesting a

LEC2-mediated negative role of this PGR , at least in this species

[

72 ]. Controversial results about ethylene-control of SE are dis-

cussed in the following paragraph.

Genetics and Physiology of Somatic Embryogenesis

58

4 The Eliciting Effects of PGRs on SE Induction and the Crosstalk with TFs

Auxin, but also other PGRs, for example cytokinins, are involved

in the speciﬁ cation and maintenance of stem cells in the zygotic

embryo and in the meristems, both in planta [ 75 – 78 ], and in

in vitro culture [

61 ]. Usually, auxin is required to induce SE in

in vitro culture, in particular the synthetic auxin 2,4-D ([

49 ], and

references therein). Moreover, auxin gradients are needed to trig-

ger the formation of stem cells in the zygotic and somatic embryo s

[

3 ]. A signiﬁ cant amount of literature on auxin biosynthesis,

metabolism, and transport, in somatic embryos shows that auxin

plays important roles, both in the induction of embryo formation

in culture, and in the subsequent elaboration of the proper mor-

phogenetic events during embryo development ([

49 ], and refer-

ences therein). However, there are also species in which a cytokinin

applied alone induces SE, e.g., in sunﬂ ower [

79 ], and species in

which a cytokinin must be combined with an auxin to induce the

process, for example, in Medicago truncatula , Vitis vinifera ,

Cyclamen persicum [

1 , 80 , 81 ]. The SE-inductive role of abscisic

acid ( ABA ) in carrot seedlings has been also reported [

82 , 83 ].

The external PGR supply has been supposed to cause local varia-

tions in the internal auxin concentration of explants, possibly trig-

gering de novo synthesis/relocation of endogenous auxin forms,

which contribute to somatic embryo induction. Moreover, 2,4-D

might act as an auxin either directly or modifying intracellular IAA

metabolism, and/or it may act as an inducer of stress-related genes

[

7 , 84 , 85 ]. For example, in the induction of SE in wheat leaf

explants, genes characteristic of a response to oxidative burst are

upregulated during the ﬁ rst hours of 2,4-D treatment [

86 ].

Moreover, as described above, 2,4-D might cause a DNA hyper-

methylation , as in Cucurbita pepo [

38 ], and induce MET1 gene

expression leading to genome reprogramming and acquisition of

SE competence.

In Arabidopsis , transcriptional regulatory networks, control-

ling stem cell population and maintenance, have been demon-

strated in planta in the shoot and root apices, and in the

procambium. Moreover, homologous TFs have been found to be

involved ([

9 , 87 ], and references therein), with activity beginning

during zygotic embryo formation. The relationship between

speciﬁ c TF-networks and PGR synthesis, transport, activity, catab-

olism is also elucidating for SE.

By the action of the ubiquitin protein ligase SCF

TIR1 , formed

by the SKP, CULLIN, F-BOX CONTAINING COMPLEX (or

SCF) and the TRANSPORT INHIBITOR RESPONSE 1 (TIR1),

the endogenous auxin is known to promote the breakdown of cer-

tain auxin/IAA (AUX/IAA) repressor proteins which, when

active, block the ARFs by forming inactive dimers. The AUX/IAA

Maria Maddalena Altamura et al.

59

inactivation allows ARFs to bind to the auxin-response elements

present on the auxin-responding genes, causing their activation.

Because AUX/IAA genes themselves are rapidly induced by auxin,

a negative-feedback loop is established with the AUX/IAA, newly

synthesized by auxin, restoring ARF-repression [

88 ]. The expres-

sion of IAA9 and IAA8 , two AUX / IAA transcriptional regula-

tors, has been observed in the SE of Cyclamen persicum and

Gossypium hirsutum [

89 , 90 ]. Recently in Arabidopsis it has been

shown that about 70 % of the AUX / IAA members display modu-

lated expression during SE, and the corresponding mutants are

impaired in somatic embryo formation [

41 ]. Taken together, it

seems evident that members of the ARF and AUX/IAA transcrip-

tion regulator/signaling families act in concert to modulate expres-

sion of auxin-responsive genes that are essential for SE. Interestingly,

in Arabidopsis and soybean, ARF5 / MONOPTEROS ( MP ) is

upregulated during SE-induction [

41 , 85 ]. In addition, during SE

in transgenic tobacco , MP is activated by LEC2 [

72 ], and Fig. 3 .

MP is a key gene in zygotic embryo patterning, affecting polar

auxin transport through activation of PIN1 auxin carrier [

91 ].

Embryo formation is impaired in vitro when auxin transport is

inhibited and MP repressed ([

41 ], and references therein). This

highlights the importance of a correct polar auxin-transport for

Fig. 3 Model of the cross talk among PGRs, stress, and TFs in SE-induction. Other proteins, exhibiting a pivotal

role in the process, are also shown. Early and further increases in endogenous IAA are shown by different

colors (details in the text)

Genetics and Physiology of Somatic Embryogenesis

60

proper SE formation, as well as for zygotic embryo formation

(Fig.

2 ). In addition, the early establishment of auxin gradient and

PIN1-mediated polar auxin transport are essential, at least in

Arabidopsis , for the induction of WUS in the callus before the mor-

phological identiﬁ cation of the somatic embryo nic cells [

92 ].

CLV3 transcripts appear later than those of WUS , and are localized

in the stem cells of the somatic proembryo [

3 ]. Like WUS role in

early deﬁ ning the shoot apical stem cell niche during SE, the WUS-

RELATED HOMEOBOX TF WOX5 is also early activated by the

auxin present in the medium, early deﬁ ning the root apical stem

cell niche, e.g. in grapevine , Arabidopsis and Medicago truncatula

SE [

64 , 93 , 94 ]. As detailed in the paragraph about Programmed

Cell Death (PCD), PCD takes part to somatic embryo develop-

ment . Polar auxin transport is essential for apical-basal patterning

and related stem niche positioning during early embryogenesis,

and disturbed transport causes aberrant embryo development, as

well as altered PCD, e.g. in Scots pine [

95 ].

It is known that auxin regulates stem cell positioning and

maintenance during plant developmental processes via an auxin

gradient resulting from a local auxin biosynthesis, coupled with

polar auxin transport ([

61 ], and references therein). A family of

YUC gene s encoding ﬂ avin mono-oxygenases, key enzymes in

auxin biosynthesis, is also required for the establishment of the

basal part of the zygotic embryo and for embryogenic organ initia-

tion. Multiple mutations of YUCs impair local auxin distribution,

resulting into severe developmental defects which resemble those

caused by multiple mutations in PIN genes [

96 , 97 ] and indicating

that auxin biosynthesis and transport are both required for zygotic

embryogenesis . This seems also the case for SE, because together

with the essential role of the auxin transport discussed before, also

YUC-mediated auxin biosynthesis has been demonstrated to occur

during SE [

4 , 69 ]. Moreover, the above-described LEC2 TF

exhibits a positive role in controlling the YUC-mediated auxin bio-

synthesis, associated with SE induction in Arabidopsis [

69 ].

Moreover, LEC2 downregulates genes involved in ethylene signal-

ing pathway in SE of transgenic tobacco [

72 ]. Interestingly ethyl-

ene disturbs SE initiation in Arabidopsis through inhibiting YUC

gene expression [

4 ]. Roles for ethylene in SE are not well under-

stood, because SE-promotive/inhibiting results have been

obtained in different species and culture systems. Ethylene is con-

sidered a stress hormone, and “stress” is a major factor in inducing

SE, as discussed in a following paragraph. Stress response can take

different forms depending on the species, the immature/mature

features of the explant tissues, and the environmental parameters

used for the culture. As discussed below, wounding and 2,4-D

application are also stress-inducers, and ethylene synthesis is rap-

idly induced in response to various stresses, including wounding

Maria Maddalena Altamura et al.

61

and auxin application [ 16 ]. In Medicago truncatula , SOMATIC

EMBRYO-RELATED FACTOR1 (MtSERF1) has been demon-

strated to be essential for SE [

98 ]. MtSERF1 encodes one of the

ERF subfamily B-3 members of the AP2/ERF TF family. Its tran-

script accumulation depends on ethylene, but also on auxin and

cytokinin, i.e. the SE-inductive PGRs in this plant [

98 ]. Its ortho-

log in Arabidopsis ( At5g61590 ) is the direct upregulated target of

AtAGL15, both positively involved in the promotion of SE in

Arabidopsis [

99 ]. In addition, the ortholog of AtAGL15 in soy-

bean (i.e. GmAGL15) upregulates genes involved in ethylene bio-

synthesis, including a 1-AMINOCYCLOPROPANE-1-CARBO

XYLIC-ACID ( ACC ) SYNTHASE ( ACS ) and an ACC OXIDASE

( ACO ) which generates ethylene from ACC, and in ethylene

response, including the TFs which are the orthologs of AtERF1

and MtSERF1 (i.e., GmSERF1 / SERF2 ). This upregulation results

into increased ethylene production and SE in soybean [

99 ].

Likewise, in Pinus sylvestris an increased content of endogenous

ethylene appears to be required for SE [

100 ]. Interestingly, none

of the ACS or ACO genes upregulated by GmAGL15 in soybean

appear to be upregulated in response to AtAGL15 accumulation in

Arabidopsis , and two putative ACO genes are repressed, hypothe-

sizing differences between AGL15 regulation in the SE of different

species [

99 ], and perhaps in ethylene levels and effects on SE. In

accordance, ethylene biosynthesis has been reported to decrease

during SE in Arabidopsis , with excessive ethylene reducing YUC

expression and disrupting local auxin distribution [

4 ]. Ethylene is

known to affect auxin transport and regulate the asymmetric distri-

bution of auxin in various plant tissues [

101 ]. Once synthesized,

ethylene is perceived by a family of receptors. COSTITUTIVE

TRIPLE RESPONSE 1 ( CTR1 ) is a receptor-interaction protein

kinase whose expression negatively regulates ethylene response

([

102 ], and references therein). In Arabidopsis , mutation at CTR1

causes costitutive ethylene signaling, and inhibition of SE initiation,

but also downregulation of most YUC genes, highlighting the pos-

sibility of a negative effect of ethylene signaling on SE through inhi-

bition of YUC expression [

4 ]. Moreover, the endogenous levels of

GA are negatively related to SE potential. The lec mutants show

increased GA levels and reduced SE [

103 ], and the exogenously

supplied GA

3 decreases tissue capacity for SE induction [ 104 ]. In

support of the inhibitory effect, several genes important for the neg-

ative regulation of GA responses display a SE-speciﬁ c upregulation,

including genes coding proteins containing the conserved amino-

acid sequence Asp - Glu - Leu - Leu - Ala , named DELLA - domain [

105 ].

Similarly the GA addition reduces SE in both nontransgenic and

transgenic (i.e., 35Spro : GmAGL15 ) soybean, and genes encoding

DELLA proteins are upregulated at some stages of SE induction

[

99 ]. Ethylene and auxin are known to impact their biosynthesis

Genetics and Physiology of Somatic Embryogenesis

62

reciprocally, and ethylene is known to act cooperatively/antagonisti-

cally with GA depending on the context [

99 ], and a central role for

AGL15 seems to exist at least during SE (Fig.

3 ).

In carrot , the application of ABA to seedlings efﬁ ciently

induces SE [

82 ] and plays an important role in the induction of

secondary SE [

106 ]. By the use of different approaches to reduce

cellular ABA levels in Nicotiana plumbaginifolia , it was demon-

strated that the ABA deﬁ ciency disturbs morphogenesis at the pre-

globular somatic embryo stage, but the effect is reverted by

exogenous ABA application [

107 ]. ABA is known to induce the

expression of LATE-EMBRYOGENESIS-ABUNDANT ( LEA )

genes in late-stage zygotic embryo s. The expression of some carrot

LEA genes is also observed during SE after treatment with ABA,

and occurs via a C - ABSCISIC ACID–INSENSITIVE3 ( ABI3 )-

mediated signal transduction [

108 ]. In Arabidopsis , the genes

coding for the EMBRYOGENIC CELL PROTEIN 31 and 63

( AtECP31 and AtECP63 ) represent the homologous to the carrot

LEA genes, and are similarly induced by ABA during SE and

equally involve an ABI3 gene expression [

109 ]. In carrot, the

endogenous levels of ABA increase in response to stress treatments,

and they are particularly high during the induction of SE in com-

parison to further embryo development al stages, suggesting the

importance of an early stress-induced accumulation of ABA for

SE-induction [

83 ]. Interestingly, LEC1 upregulates the expression

of ABI3 and FUS3 [

110 , 111 ], and all the three genes are expressed

during early microspore embryogenesis in Brassica napus [

112 ]. It

seems that ABI3 and FUS3 positively regulate each other through

a feedback loop, and the GA-ABA ratio seems to determine the

developmental mode, with low GA-ABA ratio promoting the

embryo mode of development [

112 , 113 ]. AGL15 directly con-

trols the genes encoding these TFs, in both Arabidopsis and soy-

bean [

99 , 114 ]. Thus, the roles of AGL15 are multiple, in particular

having in mind that it responds to auxin levels, but it is also capable

to repress its own expression and to activate LEC2 [

114 , 115 ]

(Fig.

3 ).

Independently of the presence/absence of cytokinin in the SE

inductive medium, the involvement of cytokinin-related TFs in SE

may be expected due to the known crosstalk between auxin and

cytokinin in the control of the respective synthesis, transport, and

signaling during morphogenesis in vitro (e.g. adventitious rhizo-

genesis, [

61 ]). In the auxin-induced embryogenic cultures of

Arabidopsis numerous cytokinin-response associated TFs are

affected, including key cytokinin regulator y genes, i.e. CYTOKININ

RESPONSE FACTORS ( CRFs ) and Arabidopsis RESPONSE

REGULATORs ( AtARRs ) [

41 ]. CRFs mediate the transcriptional

responses to cytokinin involved in the regulation of embryo and leaf

development, and function together with type-B ARRs [

116 ]. One

of these ARRs , i.e. ARR10 , is upregulated in the SE of Arabidopsis

Maria Maddalena Altamura et al.

63

[ 41 ], similarly to its homolog in Medicago truncatula , i.e. MtRR1

[

117 ]. ARR10 has been proposed to play a general role in cytokinin

signal transduction throughout the life cycle of Arabidopsis , working

redundantly with other typeB-ARRs [

118 ]. However the upregulation

of speciﬁ c TFs may also inhibit SE. It is known that the embryo-

genic competence of the callus induced from alfalfa petioles is

inhibited when kinetin is replaced by thidiazuron (TDZ) ([

5 ], and

references therein). This inhibitory effect of TDZ is associated with

the upregulation of a HD-Zip II TF, named MEDICAGO SATIVA

HOMEOBOX 1 (MSHB1) [

5 ].

Taken together, the regulatory network of TFs for cellular

reprogramming leading to SE is complex, and still far to be fully

elucidated. The mechanisms by which this regulatory network

communicate with PGRs to coordinate SE is still widely unknown,

however, a model summarizing the relationships between the main

TFs and PGRs during the reprogramming of vegetative cells for SE

induction is proposed in Fig.

3 .

5 Somatic-Receptor-Kinases Involvement in the Regulatory Network for Stem

Cell Induction and Maintenance During SE

In addition to PGRs and TFs, ligand-receptor-like kinase signaling

pathways have been revealed as crucial regulators in stem cell speci-

ﬁ cation, and an intercellular leucine-rich repeat receptor-mediated

pathway has been proposed for the maintenance of plant stem cells

([

119 , 120 ] and references therein). SERK genes form a subgroup

among the genes coding for membrane-located LEUCINE-RICH

REPEAT-RECEPTOR-LIKE KINASE proteins (LRR-RLKs),

which play important roles in plant signaling pathways ([

1 ], and

references therein). Auxin, combined or not combined with cyto-

kinin, upregulates SERK genes, depending on the species ([

121 ],

and references therein). Some SERKs are positively related to

zygotic embryo genesis , e.g., in carrot [

122 ], Arabidopsis [ 123 ],

cacao [

124 ], Medicago truncatula [ 98 ], wheat [ 125 ], as well as to

apomixis [

126 , 127 ]. The positive involvement of some SERKs in

the induction of SE has been reported for a lot of dicots and mono-

cots , e.g., carrot [

122 ], Dactylis glomerata [ 128 ], Arabidopsis

[

123 ], Medicago truncatula [ 80 ], Ocotea catharinensis [ 129 ], sun-

ﬂ ower [

79 ], cacao [ 124 ], rice [ 130 ], Citrus unshiu [ 131 ], grape-

vine [

81 ], potato [ 132 ], wheat [ 125 ], coconut [ 133 ], banana

[

134 ], maize [ 121 ], and Cyclamen persicum ([ 1 ], and references

therein). Interestingly, numerous studies suggest a role in develop-

ment, at least for speciﬁ c SERKs, broader than in SE and zygotic

embryogenesis. For example, in sunﬂ ower, a SERK gene is

expressed in both SE and shoot organogenesis [

79 ]. In Medicago

truncatula , MtSERK1 is expressed in somatic and zygotic embryo-

genesis, but also in rhizogenesis in vitro, and in all types of primary

Genetics and Physiology of Somatic Embryogenesis

64

meristems in planta [ 80 , 135 , 136 ]. In addition, AtSERK1 and

AtSERK2 redundantly control microsporogenesis in Arabidopsis

[

137 ]. All together these results suggest an involvement of speciﬁ c

SERKs in stem cell formation and maintenance in planta and

in vitro. The in vitro culture of Cyclamen persicum immature ovules

has provided useful outcomes for demonstrating SERK(s) involve-

ment in stem cell formation/maintenance, because lines forming

either organs or embryos, as well as callus lines recalcitrant to

organ/embryo formation, are available for the same cultivar and

PGR condition (Fig.

4a–c ). Using this system, Savona and cowork-

ers [

1 ] isolated two SERK genes, CpSERK1 and CpSERK2 , from

the embryogenic callus. The expression of both genes was high in

the embryogenic callus, moderate in the organogenic callus, and

null in the callus showing neither SE nor organogenesis. The

expression of both genes has been shown to start in the stem cell

clumps from which the pre-embryogenic aggregates (PEAs,

Fig. 4 Cyclamen persicum callus lines obtained under the same hormonal conditions but forming either shoots

and roots (magniﬁ ed in the inset ) ( a ), or only somatic embryo s (magniﬁ ed in the inset , arrows ) ( b ), or only cal-

lus ( c ) [see also the text, and [ 1 ] for further details]. ( d ) Encapsulated seeds of cyclamen (courtesy of B. Ruffoni

and M. Savona)

Maria Maddalena Altamura et al.

65

Fig. 5c ) and the organ meristemoids, respectively, originate.

Expression continues in the pro-embryogenic masses ( PEMs ,

Fig. 5d, m ), but progressively declines (Fig. 5d ). A similar expres-

sion pattern occurs in the organ meristemoids. In mature somatic

embryo s developing from the PEMs (Fig. 5g ), and in the shoot

and root primordia developing from the meristemoids (Fig. 5e, f ),

CpSERK1 and CpSERK2 are expressed with patterns similar to

those of the zygotic embryos (Fig. 5b ) and the primary meristems

in planta (Fig. 5a ). Thus, CpSERK1 and CpSERK2 , being

expressed in the stem cells, may be regarded as markers of pluripo-

tency. Moreover, their relation with the embryogenic potential is

of interest, because their high expression maintains the trans-

amplifying derivatives of the original stem cells (PEAs) in a plu-

ripotent condition over time, and this leads to the totipotency

necessary for somatic embryo formation [

1 ]. Consequently, even if

not peculiar of SE, the two genes may be used as markers of SE in

cyclamen, allowing the screening of the calli before the macro-

scopic expression of their fate, i.e., embryogenesis, organogenesis,

recalcitrance, with this early screening important for a large-scale

production of synthetic seeds (Fig.

4d ). The feature of

CpSERK1 / SERK2 of pluripotency/totipotency markers is in

accordance with their lack of expression during PCD occurring in

the xylogenic nodules of the organogenic calli [

1 ]. Similarly,

AtSERK1 and AtSERK2 do not seem to be involved in PCD in

Arabidopsis [

138 , 139 ]. As discussed in a following paragraph,

PCD has an important role in SE, but uncoupled with stem cells.

In Arabidopsis and other species, SERK genes form a gene

family ([

1 ], and references therein). SERKs tend to function in

pairs of redundant proteins evolutionarily organized in clades

related to either AtSERK1/2 or AtSERK3/4/5 [

140 , 141 ].

CpSERK1 and CpSERK2 are tightly evolutionarily related, and

relatively close to AtSERK1/2, and more distant to the other three

AtSERKs [

1 ]. In accordance, CpSERK1 / 2 and AtSERK1 / 2 share

a common localization in planta ([

1 ], and references therein), and

CpSERK1 and CpSERK2 are expressed in Arabidopsis , e.g. in the

root apex (Fig. 5h, k ). In addition, a pro(promoter) AtSERK1-

AtSERK1- YELLOW -FLUORESCENT-PROTEIN (YFP) con-

struct [

142 ], introduced into cyclamen embryogenic calli, expresses

correctly the fusion protein and localizes exactly as CpSERK1 /2

(Fig. 5j, c , in comparison). All together these data sustain a role for

SERK1/SERK2 complex in SE control in different species.

It is interesting to note that SERK1 and AGL15 are associated

in complexes that include components of the brassinosteroid (BR)

signaling pathway, i.e., BR-INSENSITIVE 1 (BRI1), and its core-

ceptor BRI1-ASSOCIATED RECEPTOR KINASE 1 (BAK1)/

SERK3 [

55 , 140 ]. Evidence has been presented that BAK1-LIKE

1 (BKK1)/SERK4 also participates in BR signaling [

138 ].

However, the involvement of the latter complex, as well as of BR

Genetics and Physiology of Somatic Embryogenesis

Fig. 5 Investigating SERK1/2 genes in Cyclamen persicum and Arabidopsis . ( a – g ) CpSERK1/2 RNA in situ

hybridizations on cyclamen sections. ( h , i , k , l ) expression of CpSERK1/2 in Arabidopsis , and ( j , m ) cyclamen

callus with PEMs , with expression of AtSERK1 ( j ). ( a ) Flower with strong CpSERK2 expression in stamen

67

signaling in SE control, remains to be elucidated. It might not be

excluded that SERK1 and AGL15, and perhaps LEC2, interact in

controlling SE independently on the BR pathway. In fact, it is

known that in Arabidopsis AGL15 -overexpressing tissues show

enhanced SE (see above), and have increased expression of SERK1

and reduced GA levels by the AGL15-induced GA2-oxidase ([

45 ,

53 ], and Fig. 3 ). Also in Medicago truncatula SE induction,

MtSERK1 reveals a binding recognition site for AGL15 and upreg-

ulation of GA2-oxidase [

98 ]. Moreover, upregulation of SERK1

also occurs in LEC2 transgenic tobacco in which SE is promoted

[

72 ], and a relationship between AGL15 and LEC2 in the positive

control of SE through a GA-lowering mechanism has been pro-

posed, as discussed above.

6 The Multifacet Signiﬁ cance of Stress as SE-Inducer

In vitro culture experiments have widely shown that the differenti-

ated fate of plant cells, which depends on positional information

and developmental signals in planta , can be altered under the

in vitro culture conditions, with the changes in cellular environ-

ment perceived as, and generating, signiﬁ cant stress effects. When

the stress level exceeds cellular tolerance, cells die, whereas when

the level is lower, it enhances metabolism and induces adaptation,

including gene expression reprogramming, cellular reorganization,

and developmental switch. Somatic embryo genesis may be consid-

ered among the adaptation responses to stress, because various

stresses are useful to induce this process (Fig.

3 ). For example, SE

in carrot can be stimulated by the application of heavy metal ions

(Cd

2+ , Ni

2+ , Cu

2+ , Co

2+ ), high osmotic pressure ( sucrose , NaCl)

and high temperature in the absence of exogenous PGRs ([

83 ],

and references therein). Heavy metals also induce SE in Arabidopsis

and wheat [

143 , 144 ], and high osmotic pressure or dehydration

are SE-inducers in Arabidopsis , wheat, and cotton [

143 – 145 ]. Of

course, also the excision of the explant (i.e., wounding) must be

Fig. 5 (continued) primordia and procambia. ( b ) Immature seed containing an embryo at torpedo-stage show-

ing CpSERK1 expression. ( c ) Large PEA showing uniform CpSERK1 expression. ( d ) CpSERK2 expression only

in a part of a PEM . ( e ) Adventitious root apex with CpSERK1 expression in the apical meristem and procam-

bium. ( f ) Meristematic shoot apex with strong CpSERK2 expression. ( g ) Mature somatic embryo with CpSERK1

expression in the shoot pole. ( h , k ) Whole mounts RNA hybridizations of Arabidopsis primary root apices with

CpSERK1 ( h ) and CpSERK2 ( k ). ( i , l ) Sense- probe controls. ( j , m ) Cyclamen embryogenic calli transformed with

proAtSERK1-AtSERK1-YFP construct during the induction phase . ( j ) Yellow ﬂ uorescence signal under confocal

microscopy ( arrows ) localizing AtSERK1 expression in the PEMs (courtesy of M. Savona). ( m ) Histological

control section of PEMs. See text and [ 1 ] for further details. Bars = 10 μm ( b – g ), 20 μm ( h , i , k , l ), 50 μm ( a ,

m ), 1 mm ( j )

Genetics and Physiology of Somatic Embryogenesis

68

considered as a “stressor” by a very early upregulation of the

endogenous levels of auxin, as in potato adventitious shooting

[

146 ], and of ethylene [ 147 ], but also by acting as a source of

Reactive Oxygen Species ( ROS ), as occurs in Medicago truncatula

SE [

136 ]. Moreover, ROS may induce ethylene biosynthesis [ 148 ].

It is widely known that 2-Dichlorophenoxyacetic acid (2,4-D)

may be considered as a “stressor” for the explant ([

16 ], and refer-

ences therein). For example, in soybean and potato cotyledons SE

induced by 2,4-D is associated with upregulation of oxidative

stress and defense genes [

85 , 132 ]. In agreement with a role of

ROS as stress-inducers (Fig.

3 ), SE is enhanced by increased levels

of ROS, e.g., in wheat and alfalfa [

149 , 150 ], and the application

of antioxidants to the inductive medium reduces SE in Eucalyptus

globulus [

151 ].

ABA serves as a critical messenger for stress response s, and is

considered one of the SE-inductive PGRs (Fig.

3 ). ABA increases

ROS levels in maize embryos, supporting roles of ROS in ABA-

signaling through a mechanism that still requires investigation [

16 ].

Also the micronutrient boron triggers stress-mediated path-

ways during SE, as recently reviewed [

152 ]. Boron-stress has a

direct impact on levels of ABA , e.g. increasing them as reported in

carrot SE [

153 ]. However, after ROS -signaling, a mechanism to

protect the embryogenic-potential cells against the harmful effects

of ROS is activated. GLUTATHIONE-S-TRANSFERASES

(GSTs) seem involved in this protective activity, considering the

accumulating evidence that the redox status and the glutathione

content of the cells are interrelated in plant developmental pro-

cesses. In accordance, members of the GST gene family are upreg-

ulated during auxin-induced SE of soybean [

85 ], and GST

transcripts accumulate in the somatic embryo s of numerous plants

([

49 ], and references therein).

GA has a negative effect on SE, and, in accordance, several

genes important for the negative regulation of GA show SE-speciﬁ c

upregulation in Arabidopsis , including DELLA genes [

41 ] (Fig. 3 ).

The stimulation of DELLA genes may be put in relation with the

stress response , because DELLA accumulation has been reported

to elevate the expression of genes encoding ROS detoxiﬁ cation

enzymes, reducing ROS levels [

154 ]. Moreover, results on SE in

transgenic lettuce support the hypothesis of an involvement of

SERK genes in stress-perception [

155 ].

Ca

2+ ions are key regulators of many plant developmental pro-

cesses, including sexual reproduction of gymnosperms and angio-

sperms [

156 – 158 ]. In plant cells the free ionized form of calcium

is located in the cytosol, and frequently acts as second messenger

in signaling, whereas the loosely bond calcium, which is in dynamic

equilibrium with free calcium, is present in middle lamellae and cell

wall s [

157 ]. Exogenous treatments with calcium salts have demon-

strated that calcium ions are active during organogenesis in vitro,

Maria Maddalena Altamura et al.

69

e.g., in vegetative bud formation from tobacco leaf and pith

explants, ﬂ ower and root formation from tobacco pith explants,

and root formation from Arabidopsis thin cell layer s [

159 – 161 ].

Depending on the culture system and the exogenous concentra-

tion, calcium ions affect organogenesis independently/depend-

ently of the exogenous hormone(s) [

159 , 161 ]. A positive Ca

2+ /

auxin interaction has been demonstrated in the pollen androgenesis

of Solanum carolinense [

162 ]. In the SE of carrot the process coin-

cides with signiﬁ cant variations in calcium ion distribution and lev-

els. In particular, a positive interrelation with the inductive auxin

2,4-D seems to exist because the ﬁ rst SE stages are characterized

by a strong and uniform Ca

2+ presence in the cells. Calcium-

presence, monitored by a ﬂ uorescent dye, becomes lower in PEMs ,

but again rises in the somatic embryo s from the globular to the

torpedo stage. At the latter stage, an apical-basal gradient of

calcium- distribution appears along the longitudinal axis of the

somatic embryo [

163 ]. The role of PIN efﬂ ux-carriers in the estab-

lishment of the auxin gradient, necessary to the apical-basal axis of

the Arabidopsis zygotic embryo , is well known ([

96 ], Fig. 2c ). Our

unpublished results show that calcium distribution parallels this

gradient (Fig.

2e, f ), strengthening the possible link between auxin

and calcium in axial patterning in both zygotic and somatic

embryos [

163 ]. Ca

2+ ions are also known to regulate the transcript

abundance of early auxin-inducible AUX / IAA genes [

164 ].

Moreover, in SE-induction in wheat [

165 , 166 ], Ca

2+ ions seem to

have a different role, i.e., to be involved in the induction of lipid-

transfer proteins [

86 ], which are also active compounds in SE, as

described in the following paragraph. Taken together, these evi-

dences show that there is a connection among auxin, calcium ions,

and SE induction, but this connection also involves SERKs,

because, as in wheat, SERK1 and SERK2 expression is auxin- and

calcium-dependent [

125 ]. It is important to note that calcium is

also important after SE-induction. In fact, an increase in calcium

ion supply in the medium at the time of the transfer to the auxin-

free differentiation medium, and an increased uptake by the

somatic embryos, highly enhances the number of embryos reach-

ing maturity, e.g., in carrot and sandalwood ([

7 ], and references

therein). The levels of cytosolic calcium are well known to change

transiently in response to various stresses and to auxin, as well

[

167 ], and there are evidences that this also occurs during

SE-induction, e.g., by an effect on the transduction of the auxin

signal ([

86 ], and references therein). Thus, in addition to the long-

lasting changes described above, the very rapid changes affecting

the free-calcium-pool are also of interest. For example, blocking

calcium-signaling in sandalwood cells, SE frequency decreases

[

168 ]. In this species, Ca

2+ signaling is associated with the activity

of two Ca

2+ -DEPENDENT PROTEIN KINASEs (CDPKs), and

the expression of a CDPK gene also increases during early phases

Genetics and Physiology of Somatic Embryogenesis

70

of 2,4-D induction of SE in cultured alfalfa cells [ 169 ]. In

Arabidopsis , two CDPKs have been demonstrated to activate a

stress and an ABA -inducible promoter, suggesting connection of

CDPKs to ABA-signaling pathways, with a link with calcium,

because elevation of calcium ions is sufﬁ cient to trigger ABA-

responsive gene expression [

170 ].

Nitric oxide ( NO ) is being recognized as a critical factor in

growth, development and stress response in plants [

171 ], and

there is emerging evidence that it may act as a stressor for

SE-induction. NO might exhibit its inductive role on SE affecting

the availability of Ca

2+ within the cells via protein kinases [ 172 ].

Alternatively it might increase auxin production, repressing the

basic HELIX-LOOP-HELIX protein 6 (bHLH006/MYC2), a

repressor of auxin biosynthesis, as observed in Arabidopsis SE

[

173 ]. Interestingly, NO and calcium ions are also related to PCD ,

which is essential to successful embryogenesis, as discussed in the

following paragraph.

7 Programmed Cell Death vs. Cell Survival in Somatic Embryogenesis, Two

Faces of the Same Coin

In the zygotic embryo genesis of angiosperms and gymnosperms ,

the suspensor is a terminally differentiated structure committed to

programmed cell death ( PCD ) and elimination. The suspensor

cells must die at a certain stage of embryo-proper development.

This usually occurs at the end of embryo-heart-stage, in the angio-

sperms, and at the end of the early embryogeny phase in the gym-

nosperms ([

174 ], and references therein). The suspensor is not

always formed by angiosperm somatic embryo s, but, when present

such as in carrot [

175 ] and Vitis rupestris (Fig. 6a ), its cells die via

PCD similarly to what occurs in zygotic embryogenesis. Suspensor

death by PCD also occurs in gymnosperm somatic embryos, as

described for Norway spruce ( Picea abies ) and silver ﬁ r ( Abies alba )

[

176 , 177 ]. It is important to highlight that the proper timing of

PCD in the suspensor cells is determinant to correct zygotic and

somatic embryogenesis. In Arabidopsis , mutants with altered

Fig. 6 (continued) mature somatic embryo . ( b ) Barrier formed by cells with degenerating nuclei ( arrow ) and

cutinized cell wall around a PEA of Vitis rupestris . ( c ) PEM ﬂ anked by callus cells in PCD (magniﬁ ed in the

inset ), and ( d ) mature somatic embryos of Vitis rupestris . ( e , f ) Strong Ca

2+ signal in the anther-wall tissues and

microspores of kiwifruit male-sterile anthers ( e ), and weaker Ca

2+ signal in male-fertile ones ( f ). ( g ) Nuclear

fragments showing OSMOTIN immunolocalization in a degenerating epidermal cell of an olive tree twig, also

showing a thick cuticle. ( h ) PCD nuclei of late-vacuolated microspores of kiwifruit male-sterile anthers moni-

tored by OSMOTIN immunolocalization. ( i , j ) Kiwifruit male-sterile anthers shortly before dehiscence showing

OSMOTIN-positive nuclear fragments in the middle-layer ( i , arrows ), and in the endothecium ( j , arrow ). See

text and [ 158 , 181 , 187 ] for further details. Bars = 10 μm ( a – c , e , f , i , j ), 40 μm ( d , g )

Maria Maddalena Altamura et al.

71

Fig. 6 Somatic embryo genesis from the sporophytic tissues of Vitis rupestris anthers ( a – d ), CTC-Ca 2+ signal in

kiwifruit male-sterile and male-fertile anthers ( e , f ), and PCD monitored by OSMOTIN immunolocalization in

different cell types ( g – j ). ( a ) Degenerating multilobed nuclei in the suspensor cells ( arrows ) of a Vitis rupestris

Genetics and Physiology of Somatic Embryogenesis

72

regulation of PCD in the suspensor, such as twin and raspberry ,

exhibit altered embryo development . In both mutants the suspen-

sor proliferates, and this results either into multiple embryo forma-

tion or failure of embryo transition from globular to heart stage

[

178 , 179 ]. Moreover, in Picea abies , the prolonged longevity of

suspensors delays the onset of histogenesis in the somatic embryos,

ﬁ nally resulting into disintegration of their tissues [

176 ].

Research in gymnosperms (i.e., Picea abies and Abies alba )

shows that there is another, and earlier, wave of PCD which is spe-

ciﬁ c of SE and essential for its success [

174 , 177 ]. This ﬁ rst wave

of PCD occurs in the PEMs . In Picea abies , time lapse-tracking

analysis has shown that each PEM may either multiply in the pres-

ence of the inductive PGRs (auxin and cytokinin are necessary for

SE-induction in this plant), giving rise to new PEMs, or trans-

differentiate to embryo upon withdrawal of the PGRs. The latter

pathway is only executed when massive PCD occurs in the PEM,

establishing a positive correlation between cells in PCD in the

PEM and frequency of somatic embryo s [

174 ]. This concept is

strengthened by the results of the experiments with cell-lines com-

posed of PCD-deﬁ cient PEMs, which are unable to form embryos

regardless of treatment [

180 ]. The presence of a PCD wave at the

PEM stage of angiosperm-SE still needs investigation. However, in

cyclamen SE, the progressive loss of stem trans-amplifying condi-

tion in the derivatives by the PEM couples with CpSERK1 / 2 loss

of expression (Fig. 5d ). Because the expression of these genes is

not compatible with PCD occurrence (see above), it is possible

that in the angiosperms PCD occurs as a late event in the PEM

cells, and only in those not engaged into somatic embryo forma-

tion and no more expressing SERK1 / 2 genes. In a lot of cases of

indirect SE in angiosperms, e.g., in SE from Vitis rupestris anther

tissues [

181 ], PEAs become separated from the callus cells by a

barrier of cells with a cutinized cell wall (Fig.

6b ). Events of nuclear

fragmentation leading to PCD widely occur around the encased

PEMs (Fig.

6c , and inset) and in the barrier cells (Altamura,

unpublished results). It is plausible that this out-of-PEM PCD

wave reduces the embryo-inductive potentialities in the PEMs,

giving the already existing embryogenic cells the chance to develop

further, i.e., to become a mature embryo without competition

with further forming embryonic structures. All together, in both

gymnosperms and angiosperms, PCD seems necessary to obtain a

correct SE up to the mature embryo stage (Fig.

6d ), and a PCD-

signal needs to be cell-to-cell communicated.

Zinc is a potent regulator of PCD in animals and is crucial for

correct SE patterning, as demonstrated in Picea abies SE ([

182 ],

and references therein). In this plant, high zinc accumulation in

the somatic embryo couples with a strong decrease of the ion in

the suspensor . In accordance, exposure of early embryos to a zinc

chelating agent leads to embryonic lethality, and exogenous zinc

Maria Maddalena Altamura et al.

73

supplementation suppresses suspensor terminal differentiation and

elimination, causing inhibition of embryo maturation [

182 ].

However, Zn

2+ can also have a pro-apoptotic effect on mammalian

cells [

183 ]. In accordance, in plants, when applied at high level, it

may exhibit a positive function in the PCD process. This is caused

by an increase in nitric oxide ( NO ) in the cells, which induces an

NO-mediated PCD ([

171 ], and references therein). All together

these results show that the exact level of free intracellular zinc

mediates PCD-survival decisions in embryogenesis, however other

ions seem to be also involved.

In the previous paragraph the importance of calcium in the

control of somatic and zygotic embryo genesis has been highlighted

(Fig.

2e, f ); however, the role(s) of calcium in these processes also

include a relationship with PCD . In fact, transient changes in cyto-

solic free calcium and long-lasting changes in cytosolic and cell

wall /membrane-associated Ca

2+ affect development also by induc-

ing PCD [

184 – 186 ]. Male sterility is known to occur by PCD, and

studies on microsporogenesis and microgametogenesis in numer-

ous dioecious plants have demonstrated that calcium ion distribu-

tion and content are related to this sterility ([

158 ], and references

therein). The sporophytic tissues of the anther degenerate by PCD

in both male-fertile and male-sterile plants, but PCD is delayed in

kiwifruit male-sterile genotypes, with a calcium signal in the tape-

tum, middle-layer and exine of the microspores that is higher than

in the male-fertile anthers (Fig.

6e, f ). A prolonged secretion of

calcium by the anther tissues seems to induce the inability of the

microspores to transit to microgametogenesis, causing, instead,

their PCD [

158 ].

OSMOTIN is a pathogenesis-related type-5 protein involved

in abiotic/biotic defense responses ([

187 ], and references therein).

The protein is also positively involved in PCD -induction [

188 ],

e.g., in the stem epidermal cells during cork formation (Fig.

6g ),

in degenerating sterile microspores (Fig.

6h , [ 158 ]), and in the

endosperm of olive tree seed [

189 ]. In accordance with the results

by the use of other PCD markers, OSMOTIN immunolocalization

in kiwifruit demonstrates the existence of a delay in PCD in the

sporophytic tissues of the male-sterile anthers (Fig.

6i, j ). Kiwifruit

anther tissues are able to dedifferentiate in the presence of IAA and

produce somatic embryo s, but SE only occurs from anthers of

male-fertile genotypes [

190 ]. Taken together, it seems that the

excess of calcium and the altered timing of PCD [

158 ] are posi-

tively related with the SE-inability of kiwifruit male-sterile geno-

types [

190 ]. In accordance, it is known that Ca

2+ concentration

can change the hormone-induced organogenic response. For

example, tobacco thin cell layer s were induced to produce ﬂ owers

by a speciﬁ c combination of auxin and cytokinin and a speciﬁ c

concentration of CaCl

2 , but formed vegetative buds instead of

ﬂ owers when CaCl

2 concentration was increased fourfold [ 191 ].

Genetics and Physiology of Somatic Embryogenesis

74

In addition, NO has been demonstrated to inﬂ uence the Ca

2+

availability within the cells ([

171 ], and references therein), and

NO and PCD have been shown to be involved in the stress-induced

microspore embryogenesis of barley [

192 ]. A role of OSMOTIN as

a stress- acclimating protein involved in both blocking [Ca

2+ ]

cyt

transients, and in PCD has been demonstrated in the vegetative

organs (Fig.

6g , [ 187 ]). Osmotin , and osmotin -like mRNAs have

been also found in seeds, e.g., those of Benincasa hispida [

193 ],

tobacco [

194 ] and olive tree [ 189 ], and the gene has been over-

expressed in tea and olive tree somatic embryos [

195 , 196 ]. Also

ABA , ethylene and wounding activate the osmotin gene ([

187 ],

and references therein). The transcriptionally active form NAC

[name from the ﬁ rst letters of NAM ( No Apical Meristem ), ATAF

( Arabidopsis Transcription Activation Factor ), and CUC ( Cup-

shaped Cotyledon ) genes] of AtNTL6 ( Arabidopsis thaliana N AC

with TRANSMEMBRANE MOTIF 1-LIKE) protein causes the

expression of Pathogenesis-Related-5 ( PR-5 ) genes [

197 ], and

LEC2 (Fig.

3 ) induces the expression of NAC TFs ([ 72 ], and ref-

erences therein). In Olea europaea , the NAC domain of the

homologous gene ( OeNTL6 ) induces osmotin transcription in both

seed coat and embryo, but the protein is absent in the embryo,

because of a downregulation after transcription. Concomitantly,

cuticular lipids are produced in the seed coat and extruded towards

the endosperm to enhance its cutinisation, suggesting a further

role for OSMOTIN as lipid -transfer protein [

189 ]. In accordance,

osmotin over-expression induces accumulation of oil bodies in tea

somatic embryos [

195 ]. It has been previously mentioned that

there is a phase during PEM growth characterized by the forma-

tion of a cutinized barrier around the PEMs , with PCD events

occurring in the barrier cells and in the callus around (Fig.

6b ). A

role of OSMOTIN as a lipid-transfer protein during PEM encas-

ing, and in inducing PCD around, is possible because genes

involved in encoding lipid-transfer proteins elicit PCD, e.g., in the

anther of Hordeum vulgare [

198 ], and because an OSMOTIN-like

protein accumulates and is secreted in the embryogenic cultures of

Cichorium [

199 ].

8 Hemoglobins Function as Anti-stress and Anti- PCD Compounds in SE

and Their Repression is required in the Inductive Phase

The existence of plant hemoglobins , distinct from leghemoglobin,

has been demonstrated in over 50 species ([

200 ], and references

therein). A relationship between NO , hemoglobins and PCD is

well known in mammalians, but it is also appearing in plants, e.g.,

in chicory and Arabidopsis SE [

173 , 201 ]. The main role of speciﬁ c

hemoglobins is to reduce NO levels as a result of NO scavenging,

resulting in reduction of NO toxicity and cell survival [

171 ]. Plant

Maria Maddalena Altamura et al.

75

hemoglobins have been classiﬁ ed into three groups. Class 2 hemo-

globins are upregulated by cold, cytokinin and ABA ([

171 ], and

references therein). For example, the promoter of a rice hemoglo-

bin gene is activated by ARR1 TF [

202 ], a type-B cytokinin-

responsive regulator. As discussed before, stress-induced

compounds, e.g., ROS (Fig.

3 ) and NO, are important for trigger-

ing SE, and PCD occurrence is determinant at PEM and suspensor

stages. Based on this premise, compounds causing NO-detoxiﬁ cation

and acting as anti-PCD might have a negative effect at speciﬁ c

time-points of the process. In accordance, the suppression of the

type-2 hemoglobin identiﬁ ed in Arabidopsis (GLB2/AHB2,

NON-SYMBIONTIC HEMOGLOBIN-2) enhances SE in this

species by increasing levels of NO within the embryogenic cells.

This increase causes a repression of the IAA-biosynthesis-repressor

MYC2 . Relieving the inhibition of IAA synthesis, the hormone

increases in the cells promoting WUS and SERK1 expression, and

embryogenic competence . Moreover, the repression of GLB2

increases the expression of PIN1 [

173 ], which is also essential to

SE-induction (Fig.

3 ). Similarly the suppression of Hemoglobin 1

( Hb1-2 ) gene in maize results in stimulating somatic embryo for-

mation [

171 ]. Taken together, SE is induced by compounds, such

as NO, which are known to induce PCD, but PCD does not occur

in the inductive phase because the activity of auxin, and not of anti-

PCD compounds, such as hemoglobins. Thus, a working hypoth-

esis in which SE-induction involves auxin and NO activities and

suppression of hemoglobins and PCD may be suggested. However,

the activity of hemoglobins might become essential later in the SE

process, e.g., to scavenge NO and repress PCD, at the times when

this becomes necessary.

9 Concluding Remarks

The broad repertoire of genes and complex expression patterns in

SE show that multiple cellular pathways are controlled by a con-

certed gene regulatory network. SE is an ideal model system for

investigating developmental ﬂ exibility and stem cell formation,

maintenance and polarization, in particular because there is a strict

similarity in the genetic control of zygotic and somatic embryo-

genesis in both gymnosperms and angiosperms . The ﬁ rst message

coming from the actors described in the work is that pluripotent

stem cells need to be deﬁ ned before the realization of the totipo-

tent condition. The totipotent cells result from trans-ampliﬁ cation

and maintenance over time of the original pluripotent stem cell

commitment. The switch from trans-ampliﬁ cation to trans-

differentiation (i.e., the construction of the somatic embryo) is still

obscure, but seems to share aspects with animal metamorphosis.

The lack of the switch stabilizes the cells in the embryogenic fate,

Genetics and Physiology of Somatic Embryogenesis

76

leading to secondary embryogenesis. As in animals, death ( PCD )

and life (embryo-proper) must be coordinated processes.

Differently from animals, a feedback loop exists in plants between

pluripotent and totipotent stem cells, because niches of no more

totipotent, but again pluripotent, stem cells must be present in the

mature somatic and zygotic embryo s to allow the polarized growth

of the seedling. The second message is about the importance of the

epigenome in the control of SE. Unraveling the interplay between

DNA methylation, histone modiﬁ cations , and small RNA activities

in the establishment of the epigenetic program leading to SE will

contribute to understand the behavior of plant cells in vitro and

the molecular basis of cell totipotency . A third message is about the

lipidome importance in SE, and its emerging functions in cellular

communication, trafﬁ cking control, and PCD.

The tools for biotechnology coming from these messages

are evident, because to maintain cells in a trans-amplification

state will improve the massive production of plant stem cells ,

e.g., for innovative cosmetics industry. By contrast, to stimu-

late trans- differentiation will provide a tool for improving

large-scale production of mature somatic embryo s, which is the

essential prerequisite for massive artificial seed production.

Acknowledgements

Special thanks to B. Ruffoni, M. Savona, V. De Gregis, M. Trovato,

R. Biasi, M. Kater for the contribution to our researches on somatic

and zygotic embryo genesis . Research supported by Progetti

d’Ateneo 2012, Sapienza University, Rome, Italy.

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_4, © Springer Science+Business Media New York 2016

Chapter 4

Do Mitochondria Play a Central Role in Stress-Induced

Somatic Embryogenesis?

Birgit Arnholdt-Schmitt , Carla Ragonezi , and Hélia Cardoso

Abstract

This review highlights a four-step rational for the hypothesis that mitochondria play an upstream central

role for stress-induced somatic embryogenesis (SE): (1) Initiation of SE is linked to programmed cell

death (PCD) (2) Mitochondria are crucially connected to cell death (3) SE is challenged by stress per se (4)

Mitochondria are centrally linked to plant stress response and its management. Additionally the review

provides a rough perspective for the use of mitochondrial-derived functional marker (FM) candidates to

improve SE efﬁ ciency. It is proposed to apply SE systems as phenotyping tool for identifying superior

genotypes with high general plasticity under severe plant stress conditions.

Key words Somatic embryo genesis , Mitochondria , PCD , Severe stress , Cell reprogramming ,

Phenotype plasticity , Phenotyping tool , Genotype selection

1 Introduction

It is state-of-the-art understanding that mitochondria play an

upstream role in stress response management and cell network

integration. This view is shared among scientists from various

research ﬁ elds who study organisms from diverse kingdoms, includ-

ing plants and animals [

1 – 4 ]. Mitochondria are seen central for

stress perception and orchestrating the effects of external signaling

by connecting them to growth and development control mecha-

nisms. They are movable organelles that are able to sense where

their action is required and seem to act as tuners [

2 , 3 ]. The num-

bers per cell, size, and shape of mitochondria are highly ﬂ exible

and related to development and cell function. SE is an example of

inducible developmental plasticity. It is based on molecular and

metabolic cell reprogramming that covers typically phases of dedif-

ferentiation and de novo-differentiation. SE can occur naturally

during agamospermy along a species-dependent developmental

plan and it can be induced artiﬁ cially under in vitro culture condi-

tions ([

5 , 6 ] and other chapters in this book ).

88

Changing environmental conditions is commonly referred to

as being “stressful” to organisms. This deﬁ nition of stress encloses

regular daily and seasonal, as well as unpredictable, events. These

events can have positive, negative, or even neutral effects. In cell

biology and organism physiology, stress at various levels is judged

by its effect. However, exploring stress response s is a never ending

story, since diversity is high and evolution is an ongoing process.

Nevertheless, despite this dynamics, in biotechnology and plant

breeding the view is forced to be more focused, which helps to

acknowledge that functional biology can only be understood in

case-by-case studies that respect individuality. This understanding

is getting even more important in view of the increasing awareness

of the ubiquity of endophytes and the subsequent fact that organ-

isms typically exist as complex holobionts [

7, 8 ]. Thus, the term

“individuality” must refer also to organisms that need to coordi-

nate various living units within the same body along developmen-

tal and environmental constraints. It includes not only coordinating

the diversity of cells with potentially diverse genetic, epigenetic ,

and metabolic identities, but encloses also endophytes, whenever

present. Consequently, stress effects on organism’s performance

should be assessed by upstream and superimposed measures,

because this is what will count for validating ﬁ nal stress responses.

In this context, it was proposed that plant stress responses are opti-

mization strategies dictated by thermodynamic demands, in order

to reach harmony with the environment [

9 – 11 ], and carbon bal-

ance is seen as the master integrator for plant stress responses [

11 –

13 ]. The effect of stress depends on timing, severity, and novelty of

stress events. Whether a stress will be harmful, neutral, or a driving

force for tolerance or resistance depends on the capacity and actual

engagement of the cell/tissue/organism to respond and on the

level that the organism has achieved for “optimality” in a trait-

speciﬁ c, as well as in a wider sense related to the general capacity

for phenotype plasticity [

14 ].

Nevertheless, it is possible to identify also typical stress response

patterns. In explants or cultivated plantlets exposed to successful

in vitro culture conditions for SE, the process starts typically in

distinct plant cells or group of cells. Afterwards, these cells directly

develop embryos or indirectly proliferate and form pro-

embryogenic cell structures and/or masses ( PEMs ) from which

somatic embryo s can form. Original cells and PEMs are network-

ing with their neighbors via plant common internal communica-

tion pathways, contributing to the stimulation of surrounding cells

to regain totipotency in order to run into the same developmental

direction [

6 , 15 ]. Developing embryogenic cells reproduce and

propagate the multicellular organism via a bipolar structure origi-

nating a new plantlet. These processes run similar to zygotic

embryo genesis [

15 ]. Nevertheless, the new plantlets may show

genetic differences to each other through differential genetic

Birgit Arnholdt-Schmitt et al.

89

identities in the original cells, or due to stress-induced genetic

changes ( somaclonal variation ). These changes can be expected to

result from the multiple interactions among the original cell, the

developmental stage, and the stress [

16 – 21 ]. The developmental

program for SE is heritable, and consecutively occurs in a repetitive

manner. Thus, it can be used for mass propagation. However, SE

performance can happen with varying efﬁ ciency depending on the

characteristics of individual genotypes. Low efﬁ ciency can be due

to lower strength in response and/or postponed responses. Both

can make a relevant difference for the efﬁ ciency of a SE system.

The molecular biological reasoning for differential efﬁ ciency dur-

ing both induction and initiation is still quite obscure. However,

genetic differences for efﬁ ciency in SE systems are supposed to be

mainly seen during the initiation phase [

6 , 22 ].

2 Initiation of SE Is Linked to Programmed Cell Death ( PCD )

It is now accepted that the death of cells from the suspensor and

the ﬁ nal exclusion of the suspensor itself are prerequisite to the

process of SE during bipolar patterning [

15 ]. Thus, successful SE

seems to depend on the dying of neighboring cells that initially

helped to feed the later core cells of the embryo proper. This

observation is more obvious in gymnosperms , where suspensor

structures during early and late embryogenesis are more strongly

pronounced. However, McCabe et al. [

23 ] have reported also for

carrot ( Daucus carota ) that suspensor cells die during initiation of

embryo formation via programmed cell death ( PCD ). Bozhkov

et al. [

24 ] found a two wave rhythm of PCD in Picea abies . While

one wave of PCD occurs during maturation as vacuolated PCD

and is linked to gradual degradation of the suspensor [

25 ], a ﬁ rst

wave of PCD happened already during proliferation. This ﬁ rst

period is connected to the transition of PEMs to somatic embryo s.

Smertenko and Bozhkov [

15 ] reviewed the life and death pro-

cesses during apical-basal patterning for angiosperms and gymno-

sperms both in comparison to zygotic embryo genesis . The authors

stress that the balance between survival and embryo development

and PCD together with the elimination of the suspensor are critical

for SE efﬁ ciency. Petrussa et al. [

26 ] found that in Abies alba the

rate of PCD was substantial during proliferation as well as during

the maturation stage, although much higher during proliferation.

3 Mitochondria Are Crucially Connected to Cell Death

PCD and necrotic cell death events form part of stress manage-

ment strategies for organism survival and are both related to mito-

chondrial functionality. However, while necrosis is based on

Do Mitochondria Play a Central Role in Stress-Induced Somatic Embryogenesis?

90

mitochondrial dysfunction not involved in SE observed in plants,

vacuolated PCD is connected to SE, but mitochondrial involve-

ment is more sophisticated [

27 ]. Smertenko and Bozhkov [ 15 ]

reported that mitochondria remained intact at the ﬁ nal stages of

PCD during SE, although with altered biochemical activities. The

role of mitochondria in plant PCD was described by Vianello et al.

[

28 ] and Reape et al. [ 29 ]. It was reported that mitochondrial

electrical potential and ATP levels dropped down during PCD pro-

cess [

30 , 31 ]. However, how the balance between survival and

death is established and maintained in proliferating embryogenic

cells and during the maturation phase of SE remains unclear.

Smertenko and Bozhkov [

15 ] underlined that the same groups of

protein can play a role in proliferation and cell death, depending

on their molecular environment. In maturating cells in A. alba ,

Petrussa et al. [

26 ] observed that mitochondrial activities changed

when compared to cells during the proliferation phase. The authors

found higher activity of the mitochondrial alternative oxidase

enzyme ( AOX ) in maturing cells than in proliferating cells, which

were characterized by a higher amount of dying cells. This led

them to suggest a correlation between mitochondrial activities and

the manifestation of PCD during the formation of somatic

embryo s. The alternative respiration pathway (AR) seemed to act

in the A. alba SE system as anti-apoptotic factor via reactive oxy-

gen species ( ROS ) capturing. The activities of external NADH

dehydrogenases, AOX, and the free-fatty acid circuit system were

higher in mitochondria from maturing tissues. The alternative cya-

nide-resistant pathway seemed to be activated and functional only

in maturing tissue reaching about 50 % of total O

2 uptake. It was

demonstrated a ﬁ vefold increase in this pathway compared to pro-

liferating cells [

26 ]. In contrast, the mitochondrial K + ATP chan-

nel activity was decreased, which seemed to reduce the destructive

release of cytochrome c from mitochondria. Overall, it is supposed

that mitochondria play a crucial role in the manifestation of the

two waves of PCDs during SE in conifers. A protective role of

AOX in PCD had been indicated earlier by the group of

Vanlerberghe [

32 , 33 ].

4 SE Is Challenged by Stress Per Se

It is now commonly accepted that stress induces the in vitro induc-

tion of SE ([

34 ]; reviewed by [ 5 , 6 ]). Moreover, SE is the most

pronounced example for stress-related phenotype plasticity reac-

tions [

6 ]. It is well known also that plant growth regulator s (PGRs)

are involved in wounding, plant development, and growth pro-

cesses, likewise that they are integrated in the process of external

environmental signal transmission towards the interior of organis-

mic life, and that they interfere with gene regulatory networking.

Birgit Arnholdt-Schmitt et al.

91

This is reviewed in Zavattieri et al. [ 5 ], Yang and Zhang [ 35 ],

Zeng et al. [

36 ], Osakabe et al. [ 37 ] and Fehér [ 6 ]. Several reports

show the generation of ROS or the involvement of oxidative stress

(OS) responsive genes upon SE induction conditions [

38 – 40 ].

Following a transgenic approach, Zheng and Perry [

41 ] demon-

strated that SE could be more rapid and proliﬁ c by differential

regulation of genes involved in stress response .

Appropriate abiotic stress stimuli for in vitro SE have been

empirically explored over long time, among which osmotic shock,

dehydration, water stress , heavy metal ions, pH changes, heat and

cool treatments, hypoxia, ultraviolet radiation, and mechanical or

chemical treatments, including also antibiotics ( reviewed in [

5 ]).

Several PGRs have been applied and a diversity of combinations

have been optimized not only to induce SE, but also to promote

embryo differentiation when SE is indirect and embryos are devel-

oped from a previously generated callogenic mass. However, SE in

carrot was also induced in PGR -free medium by different chemi-

cals, such as sucrose , sodium salt, or CdCl [

5 ]. SE can be regulated

by cell wall components, diverse extracellular proteins, arabinoga-

lactan protein s ( AGPs ), oligosaccharins, and through the percep-

tion and transduction of extracellular signals by receptor kinases,

Ca

2+ and its effectors, as well as by diverse transcription factor s

(reviewed in [

15 ]). Several studies show that changes in chromatin

organization and in epigenomic marks (DNA methylation, histone

posttranslational modiﬁ cations, micro RNAs) accompanies SE

induction and somatic embryo development and growth [

6 , 15 ].

These observations are not surprising, since they conﬁ rm the role

of global genome organization during normal and adaptive devel-

opment and its involvement in stress response s, also seen in other

in vitro culture systems or stress treatments [

16 , 17 , 19 ].

Nevertheless, future studies should more strongly consider differ-

ences in cell identity in the ﬁ rst responsive cells, marked not only

by differential transcript patterns [

42 , 6 ] but also by genetic and/

or epigenetic factors. This is justiﬁ ed by the current knowledge on

DNA variability at single cell and tissue levels due to single nucleo-

tide polymorphisms (SNPs), insertion/deletions (InDels), copy

number variation (CNV), and/or DNA methylation [

43 – 45 ].

Stress provided by changing environmental conditions can

promote both induction of dedifferentiation (e.g., [

46 ]) and the

realization of induced SE programs by somatic embryo develop-

ment (e.g., [

6 ]). However, due to the high diversity of inducers,

SE cannot be deﬁ ned as a speciﬁ c response to a unique stress or

stress composition. On the contrary, it must be recognized that

stress per se plays critical role as an embryonic stimulus [

5 , 6 , 47 ].

The so-called stress-induced morphogenic response (SIMR)

depends on the stress-management capacity of the plant, or of a

cell and tissue at a given developmental stage. Fehér [

6 ] pointed to

the large variation observed in SE between genotypes. He

Do Mitochondria Play a Central Role in Stress-Induced Somatic Embryogenesis?

92

highlighted also the fact that main differences among various

embryonic pathways will be found in the phase of the initiated

stress response .

5 Mitochondria Are Centrally Linked to Plant Stress Response

and Its Management

Under stress mitochondria play multiple roles. They regulate cell

homeostasis through controlling cell redox states and adapt the

supply of energy and metabolic compounds to target cell locations,

integrating stress response s with plant growth and development

both in photosynthetic and nonphotosynthetic cells [

4 , 48 ]. How

mitochondria can take over this upstream role superimposed to all

types of adaptive metabolic and morphologic cell processes related

to growth and development is currently in the focus of ambitious

research efforts and was excellently reviewed for plants by Ng et al.

[

4 ]. Crucial is the central role of mitochondria in stress perception

and transmission to cell functioning through anterograde and ret-

rograde signaling pathway networks, including dual location strat-

egies that integrate cell nucleus, cell organelles, and endoplasmic

reticulum (ER) [

2 , 4 , 49 , 50 ]. Recently, Wallace and Fan [ 51 ] and

Wallace [

52 ] highlighted in the context of human diseases the criti-

cal role of mitochondria (via bioenergetics) also for epigenetic cell

regulation.

A role of mitochondria for stress response s was conﬁ rmed for

diverse types of environmental stress stimuli that also account as

stimuli for SE. This includes importantly osmotic stress [

53 , 54 ],

salinity [

54 ], water stress [ 48 ], and temperature [ 55 ]. As reported

above some of those factors have been successful to induce SE

without any PGR application ( reviewed in [

5 ]). Sugar and hor-

mone signaling pathways interplay for the modulation of develop-

mental transition [

56 ] and it was reported that mitochondrial

invertase functions in developmental energy-demanding processes

[

12 ]. It has been shown that Glucose-TOR (Target-Of-Rapamycin)

signaling reprograms the transcriptome and activates meristems in

the control of developmental transition and growth [

13 ]. TOR

complexes constitute an ancestral signaling network, which is con-

served throughout eukaryotic evolution to control the fundamen-

tal process of cell growth. As a central controller of cell growth,

TOR plays a key role in development and aging, and has been

implicated in stress-induced disorders. This master metabolic regu-

lator was shown to be involved also in mitochondrial shaping,

which is impressively linked to mitochondrial functioning [

57 ]. It

is well known that the number of mitochondria is adaptive, depend-

ing on environmental signaling that interacts with plant develop-

ment. For example, in root cells the number of mitochondria is

plastic and correlates to induced root exudation and plant growth

Birgit Arnholdt-Schmitt et al.

93

performance. Mitochondria are highly dynamic with respect to

their biogenesis, frequent fusion and ﬁ ssion events, and size and

shape restructuring, which is related to consecutive functioning.

This dynamic seems to be regulated by tissue speciﬁ city,

developmental and internal, as well as external, stimuli [

57 – 60 ].

Vice versa, mitochondria can inﬂ uence morphogenesis as reported

for cancer [

61 ]. In plants, signiﬁ cance of mitochondria for cell fate

decisions that enclose dedifferentiation and de novo differentiation

is also recognized [

62 , 63 ]. In this context, the AR is increasingly

getting into the focus of research on stress acclimation and adapta-

tion [

64 – 66 ]. Most studies on AR focus on AOX , an inner mito-

chondrial membrane protein that functions as terminal oxidase

generating water from ubiquinol [

67 ]. The enzyme is encoded by

a nuclear gene family, which in higher plants is composed by 1–6

gene members distributed in two subfamilies ( AOX1 and AOX2 )

[

68 , 69 ]. AOX employs activity in the mitochondria at the cutting

edge of stress signal perception, cell signaling, and maintenance of

homeostasis. Several authors highlighted the involvement of also

other components of the mitochondrial energy-dissipating systems

in stress responses, such as uncoupling proteins (UCPs) and exter-

nal NADH dehydrogenases [

14 , 70 , 71 ] or other antioxidant mol-

ecules. From those, glutathione [

72 ], superoxide dismutase and

catalase [

73 ] have also been suggested as being involved in SE.

For AOX , many studies conﬁ rm a central role for cell redox

homeostasis [

74 , 75 ], a link between AOX and responses to

osmotic stress [

64 ], salinity [ 54 , 76 , 77 ] , temperature [ 70 , 78 –

80 ], drought [ 65 , 81 ], pH changes [ 82 ], nutrient limitation [ 83 ,

84 ], ozone, metal toxicity, as well as to low oxygen and high irradi-

ance ( reviewed in [

64 , 70 ]). Several reports are available also in

reference to biotic stress showing a contribution of AOX in resis-

tance against insects, virus, fungi, and pathogenic bacteria ( reviewed

in [

64 , 66 ]). AOX was proposed as “master regulator” for stress

response s [

85 ], and it is known for its involvement in the regula-

tion of seed germination [

86 ], plant growth [ 83 ] and development

[

87 ], as well in fruit development [ 88 ] and ripening [ 89 ]. The

involvement of AOX in SE was ﬁ rstly reported by Frederico et al.

[

22 ]. These authors demonstrated early differential expression of

AOX gene members during SE initiation (“realization phase”).

Application of SHAM (salicylhydroxamic acid) inhibited AOX

activities and completely suppressed embryo development .

Recently, a role for AOX in early events of dedifferentiation was

also indicated during the lag-phase of callus growth induction in

explants from carrot tap-root secondary phloem ([

90 ], Campos

et al., personal communication ). SHAM application can obviously

reveal discriminatory inhibiting effects on callus growth and devel-

opmental morphogenesis. During auxin-stimulated adventitious

rooting in microshoots from olive , SHAM application suppressed

the rooting process as expected, while simultaneously occurring

Do Mitochondria Play a Central Role in Stress-Induced Somatic Embryogenesis?

94

callus growth in the same region was not inﬂ uenced [ 91 – 93 ].

Fehér [

6 ] emphasized that calli correspond not necessarily to a

dedifferentiation state, but can also be the result of disturbed

differentiation of adult stem cells under unphysiological conditions

([

6 ], and references therein ). These results hypothesize a role for

AOX in dedifferentiation, but not a role in “misdifferentiation.”

6 Perspective View on Future Experimentation

There is no doubt that mitochondria and mitochondrial proteins

play a relevant role in plant stress response s. SE is a clear demon-

stration of the capability of plants to respond upon severe stress by

strong morphogenic plasticity in order to enable survival. Further,

SE is an example for SIMR, i.e., the stress per se is the stimulus .

Consequently, efﬁ cient biomarker and DNA marker for SE related

to applications in biotechnology or plant breeding can be supposed

to come from the mitochondrial machinery linked to cell network-

ing. This is a promising and wide ﬁ eld for future research. Since

long time it is known that induction and initiation of SE depend on

multiple interaction of [genotype × development × explant × envi-

ronment]. Recalcitrant species are well recognized ( see in this book ),

but genotype-speciﬁ c responses are also known in non-recalcitrant

species. Even within easy-to-induce species, such as D. carota , dif-

ferential responsiveness to stimuli and conditions are found at sub-

species and variety level. Based on the insight that SE is a response

upon stress, SE was proposed as a screening tool to study stress-

inducible plant plasticity as a trait per se [

14 , 94 , 95 ]. This was also

the underlying idea of initiating about 10 years ago research on

carrot SE as one of several experimental in vitro and in vivo plant

systems from diverse species, subspecies, and cultivars that show

clearly deﬁ ned developmental plasticity upon stress. In these sys-

tems the role of AOX on stress performance and its appropriate-

ness as functional marker (FM) for stress behavior was studied [

22 ,

90 , 91 , 94 , 96 – 98 ], Campos et al. ( personal communication ).

Findings from this integrated research ac ross species and systems is

expected to contribute to a better understanding of stress behavior

and phenotype plasticity as well as to advance FM development for

stress responses, including also the identiﬁ cation of markers for the

efﬁ ciency of the SE process.

The idea of Frederico et al. [

22 ] of using SE as a screening tool

for stress behavior was already taken by the breeding community.

Afuape et al. [

99 ] applied SE in cassava as a system to study stress

responsiveness of the heterologous AtAOX1a gene in search for a

linkage to post-harvest stress and its use in molecular breeding.

Similarly, a primary culture test system for carrot root explant

growth induction was used to conﬁ rm the signiﬁ cantly higher

Birgit Arnholdt-Schmitt et al.

95

responsiveness of a hybrid to yield-determining cytokinin activity

compared to growth of the according parental inbred lines [

100 ].

The system could also discriminate carrot cultivars and plant- speciﬁ c

response to temperature and reveals now AOX involvement in

dedifferentiation and growth maintenance ([

90 ], Campos et al.,

personal communication ). Recently, bioenergetics and mitochon-

drial respiration are in the focus of plant breeding research on abi-

otic and biotic stress tolerance and FM development. SE systems

might develop as important species-speciﬁ c deep phenotyping

tool , in order to screen for superior genotypes that can cope with

severe stress conditions.

Recognizing the ubiquity of endophytes in organismic life

challenges not only our fundamental understanding of functional

biology but will also drive innovation in conventional and

FM-assisted plant breeding [

8 ]. Future research needs to consider

the signiﬁ cance of plants as holobionts that should also be explored

to understand the origins of variable competence for SE. Studying

the meaning of endophytes in SE systems will be a fascinating area

of research for the coming generations of scientists. Mitochondria

are one of the most prominent examples of invasion of organisms

with mutually beneﬁ cial effects and shared coordination of the

whole organism structure and function. Improving our under-

standing of mitochondrial dynamics (variable number per cell,

mobility and, plastic sizes and shapes) related to plant morphogen-

esis will certainly contribute to improve SE efﬁ ciency.

Finally, linking bioenergetics and thus the importance of mito-

chondria for epigenome regulation may become highly instrumen-

tal for application in biotechnology and breeding. This is a research

area which can be excellently studied on SE as an experimental

system.

Acknowledgement

This work was supported by FEDER Funds through the Operational

Program for competitiveness Factors—COMPETE, and National

Funds through FCT under the Strategic Project PEst-OE/AGR/

UI0115/2014 and the project FCOMP-01- 0124-FEDER-009638

(PTDC/EBB-BIO/099268/2008). The authors are thankful to

the Portuguese FCT—Foundation for Science and Technology

(FCT) for the support given under the program POPH—Programa

Operacional Potencial Humano (Ciência 2008: C2008-UE/

ICAM/06) and also to ICAAM for the support given to H.C.

(BPD UÉvora ICAAM INCENTIVO AGR UI0115) and C.R.

(BTI_Uevora_ICAAM_PTDC_EBB-BIO_99268_2008).

Do Mitochondria Play a Central Role in Stress-Induced Somatic Embryogenesis?

96

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_5, © Springer Science+Business Media New York 2016

Chapter 5

Dying with Style: Death Decision in Plant Embryogenesis

Shuanglong Huang , Mohamed M. Mira , and Claudio Stasolla

Abstract

Embryogenesis is a fascinating event during the plant life cycle encompassing several steps whereby the

zygote develops into a fully developed embryo which, in angiosperms, is composed of an axis separating

the apical meristems, and two cotyledons. Recapitulation of embryogenesis can also occur in vitro through

somatic embryogenesis, where somatic cells are induced to form embryos, and androgenesis, in which

embryos originate from immature male gametophytes. Besides cell division and differentiation, embryo

patterning in vivo and in vitro requires the dismantling and selective elimination of cells and tissues via

programmed cell death (PCD). While the manifestation of the death program has long been acknowl-

edged in vivo, especially in relation to the elimination of the suspensor during the late phases of embryo

development, PCD during in vitro embryogenesis has only been described in more recent years.

Independent studies using the gymnosperm Norway spruce and the angiosperm maize have shown that

the death program is crucial for the proper formation and further development of immature somatic

embryos. This chapter summarizes the recent advances in the ﬁ eld of PCD during embryogenesis and

proposes novel regulatory mechanisms activating the death program in plants.

Key words Androgenesis , Embryogenesis , Hemoglobins , Programmed cell death , Somatic

embryo genesis

1 Programmed Cell Death in Plant Growth and Development

The term programmed cell death ( PCD ) encompasses several dis-

tinct pathways unique to eukaryotes [

1 ] which lead to the selective

dismantling and elimination of cells, tissues, and/or organs. This

active process, which together with cell division and differentiation

contributes to the “shaping” of the organism, is controlled by

endogenous factors and relies on energy-dependent events [

2 ].

Manifestation of PCD in plants is developmentally and environ-

mentally regulated and observable throughout the life cycle. The

most dramatic examples of development-regulated PCD are appar-

ent during xylogenesis, the maturation and death of xylem cells

required for the formation of the vascular system, reproduction,

involving the elimination of speciﬁ c embryogenic cells or the selec-

tive killing of female primordia, and senescence, where PCD

102

ensures the removal of old tissue and the turnover of macromole-

cules [

3 ]. Activation of PCD is also triggered by some abiotic and

biotic stresses. While during ﬂ ooding conditions selective removal

of cortical cells maintains a continuous supply of oxygen to the

under-water organs through the formation of aerenchyma, during

plant-pathogen interaction programmed elimination of cells limits

pathogen growth and reduces the infection sites [

3 ].

In animal systems, the mechanisms regulating PCD have been

well investigated and, based on morphological, biochemical and

molecular characteristics, three types of PCD are distinguished:

apoptosis, autophagy, and necrosis [

1 ]. In plants, despite the early

recognition of PCD, knowledge on the biochemical and molecular

events underlying PCD is scarce and classiﬁ cation of the death

pathways is solely based on morphological criteria [

4 ]. According

to van Doorn [

5 ], PCD in plant cells can be categorized as necrosis

and vacuolar cell death. While the former is generally caused by the

rupture of the plasma membrane and the shrinkage of the cytoplas-

mic components, as often observed under abiotic stresses, the lat-

ter is characterized by the clearance of the cytoplasm triggered by

the rupture of the tonoplast and the release of vacuolar hydrolytic

enzymes. Vacuolar cell death is very common during development

where it is involved in organ formation. It must be kept in mind

that the categorization of cell death into these two pathways, i.e.,

necrosis and vacuolar cell death, is somehow simplistic as some

atypical examples of cell death do not follow in either category [

5 ].

2 Ultrastructural and Cytological Characteristics of Necrotic and Vacuolar Cell

Death in Plants

Considered for a long time an “unprogrammed” event, necrosis

has been recently included as an integral pathway of PCD [

5 ] char-

acterized by two early hallmarks: the increase in cellular volume

and the rupture of the cytoplasm leading to the release of the intra-

cellular content [

1 ]. Although poorly characterized in plants,

necrosis in animals is also accompanied by increases in cytosolic

Ca

2+ and changes in mitochondrial and lysosomal function leading

to the accumulation of reactive oxygen species ( ROS ) [

6 ]. As sum-

marized by van Doorn [

5 ], necrosis is typical of the hypersensitive

response and cells challenged with necrotrophic pathogens.

Unlike necrosis, vacuolar cell death is better characterized and

manifested by the rupture of the tonoplast and the release of the

hydrolytic enzymes. Plant cells are equipped with two major types

of vacuoles: storage vacuoles which accumulate preferentially pro-

teins, and lytic vacuoles enriched with several hydrolytic enzymes

including aspartate and cysteine proteases and nucleases [

5 ].

Manifestation of vacuolar cell death can be non-disruptive, if the

tonoplast fuses with the plasma membrane and releases the

Shuanglong Huang et al.

103

hydrolytic enzymes in the apoplast, or disruptive, if the collapse of

the tonoplast discharges the hydrolytic enzymes within the cyto-

plasm [

7 ]. This second series of events has been shown to occur

during lysogenous aerenchyma formation through three tempo-

rally distinct steps. The ﬁ rst step is characterized by the swelling of

the lytic vacuoles which occupy most of the symplast. During the

second step, the tonoplast invaginates and through processes anal-

ogous to autophagy of animal cells engulfs and degrades cytoplas-

mic regions [

8 ]. Microscopy studies revealed shrinkage of the

plasma membrane and the formation of granular bodies within the

lytic vacuoles and around the organelles engulfed by the tonoplasts

[

9 ]. The third and ﬁ nal step is characterized by the lysis of the

tonoplast and the release of the hydrolytic enzymes which clear

cytoplasmic components starting with the endoplasmic reticulum

and terminating with the nucleus and mitochondria [

5 ]. Deviations

from this sequence, such as the early disruption of the cell wall

prior to the rupture of the vacuole [

10 ], are observed and conﬁ rm

the simplistic classiﬁ cation of the proposed death pathways.

The most characteristic cytological events of PCD are visible

in the nucleus and compromise the ability of the DNA to tran-

scribe and replicate; these include the degradation of DNA, the

condensation of chromatin, and nuclear fragmentation [

11 ].

Degradation of DNA is executed by nucleases and occurs in two

distinct phases: the initial cleavage of the DNA at the interloop

sites of the chromatin producing DNA fragments of about

50–300 kbp, followed by cleavage at the internucleosomal sites

which generate 200 bp DNA fragments [

12 ]. These events occur

in conjunction with the condensation of chromatin which

requires de-polymerization of F-actin [

13 ], and the fragmenta-

tion of the nucleus which is very typical of animal apoptosis [

14 ].

Although nuclear fragmentation is generally one of the last events

of PCD, it was reported as the ﬁ rst sign of PCD during aerenchyma

formation in oxygen -deprived plants [

15 ].

3 Execution of PCD During Plant Embryogenesis

Embryogenesis is an important event during the plant’s life cycle.

The zygote, originating from a single fertilization event in gymno-

sperms and a double-fertilization event in angiosperms , undergoes

a precise pattern of cell divisions culminating in the formation of a

fully developed embryo. The subsequent imposition of a matura-

tion period, in which the seed experiences water stress , is required

for the termination of the developmental program and the initia-

tion of germination [

16 ]. Recapitulation of embryogenesis can

also be achieved in culture through judicious manipulations of

media and culture environment. Two methods routinely used to

generate in vitro embryos are somatic and gametophytic

Cell Death in Plant Embryogenesis

104

embryogenesis. While the former method is employed to generate

embryos from somatic cells, i.e., cells other than gametes, the latter

uses male or female gametophytes as explants. The utilization of

male gametophytes to produce embryos ( microspore embryogen-

esis , sometimes referred as androgenesis ) exploits the ability to re-

route the developmental fate of immature pollen , i.e., microspores,

from a gametophytic to an embryogenic pathway [

17 ]. Both

somatic and gametophytic embryogenic systems are used as model

systems to investigate biochemical and molecular events governing

embryo development .

Execution of PCD is an integral component of embryonic

development both in vivo and in vitro as it shapes the body of the

embryo through the elimination of speciﬁ c cells and organs.

Experimental interference with the death program compromises

the formation of the embryos [

18 ].

Manifestation of PCD is apparent during different phases of in vivo

embryogenesis. It participates in the dismantling of the suspensor ,

the removal of supernumerary embryos produced by polyembry-

onic seeds, and degradation of nucellus , endosperm , and aleurone

layer. This chapter only deals with the ﬁ rst two events as they are

intimately related to the formation of embryos. Detailed descrip-

tions of the last events are available [

19 ].

Formation of the suspensor is concomitant to that of the embryo

proper. In angiosperms , the ﬁ rst asymmetric division of the zygote

originates an apical cell and a sub-apical cell. While the apical cell

gives rise to the embryo proper (which will progress though a

globular, heart, cotyledon, and torpedo stage of development),

transverse divisions of the subapical cells generate the suspensor

[

20 ]. Besides its passive function in anchoring the embryo to the

seed, the suspensor plays two key roles. It transfers nutrients to the

embryo proper and it participates in the establishment of the polar-

basal embryonic axis by modulating the ﬂ ow of auxin [

21 ]. The

suspensor is short-lived and once its functions are no longer

needed, generally at the cotyledon stage of development, it is dis-

mantled through the execution of the death program [

22 ]. In all

cases examined, PCD is required for the elimination of the suspen-

sor regardless of its shape and morphology which differ remarkably

among species. While in orchids the suspensor consists of a single

cell, the Arabidopsis suspensor is composed of about seven cells

while runner bean suspensors have more than 200 cells [

23 ].

Variations in the number of suspensor cells are also observed within

the same family [

24 ]. Profound differences in suspensor morphol-

ogy are also apparent. In angiosperms, the suspensor is generally

composed by a ﬁ le of single cells characterized by two regions: the

neck including suspensor cells adjacent to the embryo proper and

the knob comprising suspensor cells in close proximity to the seed

3.1 Role of PCD

During In Vivo Plant

Embryogenesis

3.1.1 Elimination

of the Suspensor

Shuanglong Huang et al.

105

integuments [ 25 ]. More complex morphological arrangements are

observed in gymnosperms , such as in Picea abies , where the sus-

pensor consists of deﬁ ned tiers of cells with the upper tier “embry-

onal tube cells” produced by the asymmetric division of the embryo

proper [

22 ]. Independent evidence suggests that elimination of

the suspensor by PCD progresses basipetally, starting from the top

suspensor cells adjacent to the embryo proper and terminating to

the bottom portion of the suspensor. Using Phaseolus coccineus as

a model system, Lombardi et al. [

25 ] showed the basipetal spread-

ing of DNA fragmentation, a hallmark of PCD, from the neck

region (top) to the knob region (bottom) of the suspensor. This

“death” pattern was also observed in maize [

26 ] and in spruce

[

22 ]. Contrasting reports describing an acropetal movement of

PCD in suspensor cells exist, but they are solely based on ultra-

structural evidences and are not substantiated by PCD marker

analyses [

27 ]. As reviewed by Bozhkov et al. [ 22 ] two scenarios

have been proposed to account for the progressive development of

PCD. The ﬁ rst involves the presence of a “cell-death” signal pro-

duced by the embryo proper which is released basipetally towards

the suspensor cells, while the second would require the depletion

of an “anti-death factor.” The generation and analyses of suspensor

mutants might resolve the nature of the PCD progression. An

intriguing question arising from the progressive spreading of PCD

is whether the suspensor cells are committed to die only after they

are fully differentiated. Arabidopsis suspensor cells are targeted by

PCD only after the suspensor is fully formed, thus suggesting that

death occurs in terminally differentiated cells. This notion is also

substantiated by analyses of tween mutant embryos. In these

mutants, suspensors cells can re-differentiate into embryogenic

cells and this ability is retained only up to the globular stage, after

which cells become fully differentiated and committed to die [

28 ].

In spruce, however, the death program is initiated in newly formed

suspensor cells which are not terminally differentiated. Suspensor

cells are added from asymmetric divisions of the embryo proper

and they start dying soon after they are formed [

22 ].

Elimination of the suspensor cells is a slow process and cells

are not subjected to rapid disruption. This is possibly required for

the proper differentiation of the embryo, given the function of

the suspensor in transporting nutrients [

22 ]. Time-course analy-

ses of zygotic and somatic embryo genesis in gymnosperms [

29 ,

30 ] suggest that the death and removal of a suspensor cell occurs

in a period of at least 5 days [

22 ]. While the information above

argues strongly for the involvement of PCD in the elimination of

the suspensor, it provides a rather simplistic picture on the timing

and progression of PCD, as variations in both are apparent.

Different patterns of PCD timing and progression in suspensor

cells exist and, in the case of the Leguminosae family, have been

categorized [

31 ].

Cell Death in Plant Embryogenesis

106

Generation of multiple embryos from one zygote is referred to as

monozygotic polyembryony, a common event in animal reproduc-

tion [

32 ]. This process, the genetic bases of which are unknown,

requires the physical splitting of cells after a few rounds of mitotic

divisions leading to the formation of two or more genetically iden-

tical embryos. Although several plant species develop supernumer-

ary embryos, monozygotic polyembryony is particularly common

in gymnosperms where several embryos are produced in one seed,

but only a “dominant” one continues to complete the develop-

mental process. The remaining “subordinate” embryos are elimi-

nated [

33 ]. Based on the growth rate of the supernumerary

embryos, Filonova et al. [

34 ] divided the development of pine

seeds into three distinct phases. The ﬁ rst phase is characterized by

the formation of multiple embryos from the same zygote. The

embryos share the same growth rate with no dominance.

Acquisition of “dominant” characteristics of one embryo, which

overgrows the subordinate embryos, demarks the second phase. In

the third phases, the subordinate embryos are eliminated by PCD ,

while the dominant embryo completes the maturation process.

Manifestation of PCD in the subordinate embryos follows a spe-

ciﬁ c pattern and is part of two distinct death programs [

22 ]. By

using terminal deoxynucleotydil transferase (TdT)-mediated

dUTP nick-end labeling (TUNEL) to detect DNA degradation in

the subordinate embryos, Filonova et al. [

34 ] showed that PCD is

initiated in the basal part of the embryo proper and progresses

acropetally reaching the apical cells in approximately 4 weeks. As

reviewed by Bozhkov et al. [

22 ], this pattern can be established by

the presence of a “death-inducing signal” moving acropetally, or

by a survival signal which accumulates preferentially in the apical

region of the embryo. The nature and origin of the signal are

unknown, but the signal might be produced by the megagameto-

phyte where PCD precedes the autodestruction of the embryonic

cells [

34 ]. This idea is also resonated by Young and Gallie [ 35 ]

who proposed that death in the megagametophyte precludes the

transport of nutrient (and a putative survival signal) to the super-

numerary embryos. Orchestration of the death program in mega-

gametophytic and embryonic cells is in fact required for normal

seed development [

22 ]. As in any other developmental process

governed by PCD, the response to the death program can be quite

ﬂ exible and environmentally inﬂ uenced, as demonstrated by the

presence of more than one dominant embryo able to develop to

maturity [

36 ].

As outlined above, recapitulation of the embryogenic process can also

occur in vitro through somatic and microspore -derived embryogen-

esis. In both processes removal of cells through PCD is an integral

component of embryo development , and recent studies have emerged

on the molecular components governing the death program.

3.1.2 Monozygotic

Polyembryony: Survival

of a Single Dominant

Embryo

3.2 Role of PCD

During In Vitro Plant

Embryogenesis

Shuanglong Huang et al.

107

While descriptions of hormone- and density-induced death pro-

grams have been shown in several somatic embryo genic systems

[

37 – 39 ], it was von Arnold’s group to ﬁ rst prove the requirement

of PCD for proper spruce embryo development in vitro [

30 , 34 ].

The spruce system is suitable for these studies as its developmental

pathway has been well characterized [

30 ]. In this system the pro-

embryogenic masses ( PEMs ), generated from cultured zygotic

embryo s, are maintained with the plant growth regulator s auxin

and cytokinin (PGRs), and consist of three deﬁ ned cellular aggre-

gates ( PEM I-III). Proembryogenic masses I (PEM I) are com-

posed of clusters of highly cytoplasmic cells subtended by a single

suspensor -like cell . Addition of other suspensor cells to PEM I

forms PEM II. As more cells are added to the PEM II the cluster

grows in size and differentiates into PEM III. In the presence of

PGRs, the three PEMs co-exist without forming embryos. Removal

of PGRs from the culture medium stimulates the trans-

differentiation of PEM III into somatic embryos through processes

involving PCD [

30 ]. The massive execution of the death program

in the PEM III re-shapes the cell cluster and allows the formation

of somatic embryos. Independent evidence suggests that PCD is

obligatory for proper and successful embryogenesis. Besides the

positive correlation observed between the extent of PCD in the

PEM III and the number of somatic embryos produced [

30 ], inhi-

bition of PCD through manipulations of the culture medium com-

promises the differentiation of PEM III into somatic embryos

[

18 ]. Consistent with the requirement of the death program, lines

composed by PCD-deﬁ cient PEMs are not able to form embryos

[

40 ], possibly because of their inability to reprogram their tran-

scriptional machinery [

41 ]. These results are analogous to

Drosophila studies showing that blockage of PCD by mutagenesis

results in prenatal death [

42 ].

Ablation of the PEMs III by the death program is followed by

a second wave of PCD which removes the suspensor of the somatic

embryo s during the late phases of development. This second wave

follows a basipetal gradient, starting from the suspensor cells adja-

cent to the embryo proper and proceeding towards the basal region

of the suspensor [

43 ].

Microspore -derived embryogenesis relies on the ability of imma-

ture microspores to redirect their normal gametophytic develop-

mental pathway toward a sporophytic route. This cell

reprogramming can be triggered in culture by imposition of diverse

stress treatments including heat shock, starvation, cold conditions,

and ethanol and gamma irradiation [

44 ]. According to Touraev

et al. [

17 ] the embryogenic process involves two steps: the forma-

tion of multicellular structures (MCS) within the exine wall of the

isolated microspores, and the differentiation of MCSs into embryo-

like structures (ELS). Formation of MCSs can occur through

3.2.1 PCD During

Somatic Embryogenesis

3.2.2 PCD During

Microspore- Derived

Embryogenesis

Cell Death in Plant Embryogenesis

108

different pathways. In the ﬁ rst pathway, the microspore nucleus

divides asymmetrically forming a generative and a vegetative cell.

Divisions of the vegetative cells give rise to MCSs [

45 ]. In the sec-

ond pathway, common to rapeseed, potato and tobacco , MCSs are

formed directly from symmetric divisions of the microspore

nucleus. The ﬁ rst pathway is characterized by the simultaneous

divisions of the vegetative and generative cells, both contributing

to the formation of MCSs [

46 ].

Manifestation of PCD is an integral component of microspore-

derived embryogenesis, especially during the early phases, as most

of the anther tissue harboring the microspores undergoes massive

death. Degeneration of cells by PCD is ﬁ rst apparent in the tape-

tum of the anthers at the pre-meiosis stage [

47 ]. As reviewed by

Varnier et al. [

48 ], this ﬁ rst death wave, which contributes to the

total elimination of the tapetal cell layer, has a temporary effect on

the microspores soon after meiosis, as some death “information”

might migrate from the dying tapetum to the microspores. Signs

of microspore degradation are often observed [

49 ] and this might

compromise their redirection towards the embryogenic pathway.

Therefore, the ability to control and manipulate the course of the

PCD process in the microspores is crucial for ensuring a high

recovery of embryos. Does the stress pretreatment, which redirects

the developmental fate of the microspores towards the embryo-

genic pathway, interfere with the death program? Wang et al. [

50 ]

showed that while inducing death in the tapetal cells, the stress

pretreatment does not accelerate death in the microspores.

Furthermore, at a metabolic level components of the PCD machin-

ery, including the bax inhibitor Bi1, are induced during the stress

pretreatments [

51 ]. Based on the above results, Varnier et al. [ 47 ]

suggested that the arrest of the death pathway in the microspores

is a necessary prerequisite for redirecting their fate towards embryo-

genesis. Once the redirection step has occurred the microspores

undergo a symmetric division and no evidence of PCD is apparent,

as revealed by transcriptome and proteomic studies [

52 , 53 ].

Contrasting observations were reported in barley , where micro-

spores subjected to stress pre-treatment exhibited increasing levels

of cell death [

54 ]. Discrepancies in results are possibly due to dif-

ferent systems and stress conditions utilized.

The second wave of PCD during microspore -derived embryo-

genesis occurs during the differentiation of MCSs into ELSs. The

development of mannitol -stressed barley microspores into haploid

embryos is characterized by formation of MCSs composed by two

distinct cell domains derived from proliferation of the vegetative

cell and generative cell, respectively. These two domains have dif-

ferent fates; the generative domain is eliminated by PCD, while the

vegetative domain develops into ELS [

55 ]. According to the

authors, the elimination of the generative domain marks the site of

exine rupture from where the globular embryos will emerge.

Shuanglong Huang et al.

109

Collectively, these studies suggest that PCD plays an integral

and important role during (1) the redirection of the microspores

from a gametophytic to an embryogenic pathway, and (2) the early

morphogenetic events associated to embryo development . The

capacity to experimentally manipulate the death program during

both processes would provide valuable insights into the require-

ment of PCD for microspore -derived embryogenesis.

4 Regulation of PCD During Embryogenesis

Regulation of the death program in plants is complex and relies on

the participation of many components, some of which participate

in unrelated responses. A premise to any investigation on plant

PCD should be that the death program in plants is mediated by

factors fulﬁ lling similar functions to regulators of animal PCD. The

following section outlines the role of some proteins and signal

molecules in modulating the cell survival/death decision during

embryogenesis.

Animal apoptosis is mainly orchestrated by the Bcl-2 related pro-

teins, which include pro-survival and pro-apoptotic members.

While pro-apoptotic members, such as those of the Bax subfamily,

trigger death events through the release of cytochrome C from the

mitochondria , pro-survival components, such as Bax-inhibitor 1

( BI-1 ) abrogate these events [

56 ]. Initially characterized in humans

for its ability to repress the yeast death pathway activated by the

over-expression of the mouse Bax gene [

57 ], BI-1 has been iso-

lated in many species of yeasts, plants and animals where it is

expressed under stress conditions and in senescent tissues [

58 , 59 ].

The pro-survival nature of this protein was also deﬁ ned in plants

through transformation studies. Cell death induced by pathogens,

fungal elicitors, temperature stress and hydrogen peroxide was

repressed in Arabidopsis plants ectopically expressing BI - 1 [

60 ]. In

the same line, a down-regulation of BI-1 accelerated death in car-

bon starved tobacco cells [

61 ]. Although the role of BI-1 has not

been investigated during embryogenesis, Maraschin et al. [

51 ]

showed a transcriptional activation of the barley BI-1 following

stress treatments which induce embryogenesis possibly through

the suppression the PCD pathway. Localization studies together

with analyses of structural and functional domains suggest BI-1

proteins reside in the ER membranes where they have a protective

role against the ER-stress induced PCD, a condition where basic

ER functions are compromised [

62 ]. This cytoprotective role of

BI-1 is mediated by its ability to modulate Ca

2+ homeostasis and

response in the ER by interacting with several calcium-binding

proteins [

63 ].

4.1 Bax-Inhibitor-1

Cell Death in Plant Embryogenesis

110

As indicated above, the pro-survival role of BI-1 proteins is to

counteract the pro-death effect of other factors, including kiss of

death, a small amino acid peptide which triggers PCD [

64 ]. Using

two mutant kod alleles and KOD over-expressing lines, the authors

demonstrated the involvement of KOD in the elimination of the

Arabidopsis suspensor during embryogenesis and its participation

in early PCD events including the depolarization of mitochondrial

membrane [

64 ]. The ability to trigger the suicide program by the

sole expression of KOD makes this gene a suitable tool to target

and dismantle cells by PCD.

Apoptosis in animal cells is triggered by the activation of caspases,

proteolytic enzymes able to cleave proteins at speciﬁ c amino acid

residues. Expressed as inactive pro-enzymes, i.e., pro-caspases, cas-

pases are activated at the onset of the death program where they

initiate an irreversible proteolytic cascade of events involving the

induction of other caspases and culminating to rapid cell death

[

65 ]. Based on their position and function along this cascade, cas-

pases are broadly divided into initiators (caspase 2, 8, 9, and 10),

executioners (caspase 3, 6, and 7) and inﬂ ammatory (caspase 1, 4,

and 5) [

66 , 67 ].

While direct homologues of caspases are not found in plants,

proteins with similar functions have been identiﬁ ed as metacas-

pases , characterized by caspase-like secondary structures and cata-

lytic domains [

68 ]. Involvement of metacaspases in PCD has been

demonstrated in yeast, where the survival/death fate is dependent

upon metacaspase expression [

69 ], and more recently in plant

embryos [

70 ]. In this latter study, it was showed that a spruce

metacaspase (mcII-Pa) is expressed during spruce somatic embryo-

genesis in tissues committed to PCD, i.e. suspensor of immature

embryos and procambium of late embryos, and that RNAi-

mediated suppression of mcII - Pa prevents the differentiation of

somatic embryos from PEMs III by repressing the death program.

Besides establishing metacaspases, and mcII-Pa speciﬁ cally, as exe-

cutioners of PCD in plant embryogenesis, this work emphasized

the relevance of the death program for the formation of embryos

in culture. The cellular function exercised by mcII-Pa requires its

cysteine-dependent arginine-speciﬁ c proteolytic activity and its

ability to migrate from the cytoplasm into the nucleus to induce

the fragmentation of DNA and the disassembly of the nuclear

envelope in cells committed to die [

71 ].

Nitric oxide ( NO ) is a signal molecule fundamental for a broad

range of plant developmental and environmental responses, includ-

ing hormone signaling, cell cycle mechanisms, and biotic and abi-

otic stress response s [

72 ]. Over the past years its role as modulator

of PCD has emerged and NO participation during the embryo-

genic process has received increasing attention. In animals, the

4.2 Metacaspases

4.3 Nitric Oxide

Shuanglong Huang et al.

111

pro-apoptotic role of NO occurs through several mechanisms.

Besides inducing two caspase activators: p53 and CD95 [

73 ], NO

inﬂ uences the death program by modulating protein nitrosation/

nitrosylation and the level of cellular cGMP [

74 ]. During plant

pathogen interaction, protein nitrosylation via reaction with NO

regulates the activity of many stress-related enzymes, including

metacaspase 9 [

75 ]. Nitric oxide also inﬂ uences the pool of cGMP

by binding to the ferrous heme group of the guanylate cyclase-

coupled receptor converting GTP to cGMP, an effector of apopto-

sis [

76 ]. In plants, administration of NO increases PCD through

an elevation of cGMP which opens Ca

2+ channels through inter-

mediates including cyclicADP-ribose [

77 ]. A spike in cellular Ca

2+

increases mitochondria permeability and triggers the death pro-

gram [

78 ]. Of note, applications of 8-Br-cGMP, a cGMP analog,

suppress caspase activity and PCD [

79 ]. While these regulatory

mechanisms have not been demonstrated during embryogenesis,

the NO-mediated activation of caspase activity and PCD has been

recently shown to occur during the early phases of microspore -

derived embryogenesis [

54 ].

As suggested above, NO homeostasis is crucial for cell death/

survival decision and plant hemoglobins ( Hbs ) are active NO scav-

engers [

74 ]. Plant HBs have been classiﬁ ed into three classes

depending on their structural and chemical characteristics, but all

of them react with NO producing nitrate and oxidizing ferrous

hemoglobin to methemoglobin [

72 ]. Studies in animal systems

have shown the ability of Hbs to inﬂ uence the death program by

modulating NO, a function that we have shown to be retained

during plant embryogenesis [

80 , 81 ]. Suppression of two Hbs

( ZmHb1 and ZmHb2 ) in maize embryogenic tissue induces PCD

by increasing NO levels in cells in which Hbs are repressed. This

increase of NO produces opposite outcomes on embryo yield

depending on the expression patterns of the two Hbs . While sup-

pression of ZmHb1 , which is expressed in both suspensor cells and

embryo proper, triggers massive death resulting in embryo abor-

tion, suppression of ZmHb2 , which is expressed solely in a few cells

anchoring the embryos to the embryogenic tissue, eliminates these

“anchor” cells releasing the embryos in the culture medium,

encouraging their growth, and increasing total embryo production

[

81 ]. The induction of PCD in Hb -suppressing cells ﬁ ts a model in

which repression of Hbs causes localized NO maxima which

increase intracellular Zn

2+ levels, by favoring its release from

metallothioneins through the destruction of the zinc-sulphur clus-

ters [

81 , 82 ]. Changes in cellular Zn

2+ homeostasis inﬂ uence the

death/survival decision in a system-dependent fashion. While in

some embryogenic systems the PCD program is induced by deple-

tion of Zn

2+ level [ 83 ], in others, including maize, an elevation in

Zn

2+ level triggers cell suicide through the MAPK cascade which

activates NADPH oxidase and induces production of reactive

Cell Death in Plant Embryogenesis

112

oxygen species [ 81 ]. Based on these observations, the authors

identiﬁ ed Hbs as potential regulators of in vitro embryogenesis by

elevating NO levels and promoting the suicide program.

5 Conclusions

Removal of unwanted plant cells by PCD is an important factor for

embryonic and post-embryonic development. During in vivo

embryogenesis activation of the death program is required for the

elimination of the suspensor , once the function of this organ is not

needed, and for the selective elimination of supernumerary

embryos in polyembryonic seeds. Recent advances on the role of

PCD during in vitro embryogenesis have evidenced the death pro-

gram as an obligatory event for the early phases of embryo forma-

tion. While the ability to alter the embryonic death program with

molecular and pharmacological approaches has been pivotal in the

identiﬁ cation of some executors of the death pathway, more infor-

mation is required to identify the early inductive signals triggering

death. Speciﬁ c attention should be addressed on the initial steps of

the death commitment, the reversibility of the commitment pro-

cess, and most importantly the identiﬁ cation of cues (positional?)

responsible for selective death occurring within the cultured tissue.

Answers to these questions will open new avenues for targeted

applications and manipulations of PCD to enhance embryo quality

and yield.

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Cell Death in Plant Embryogenesis

117

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_6, © Springer Science+Business Media New York 2016

Chapter 6

Somatic Embryogenesis in Broad-Leaf Woody

Plants: What We Can Learn from Proteomics

Sandra I. Correia , Ana C. Alves , Paula Veríssimo ,

and Jorge M. Canhoto

Abstract

Proteomic approaches have been used to understand several regulatory aspects of plant development.

Somatic embryogenesis is one of those developmental pathways that have beneﬁ ciated from the integra-

tion of proteomics data to the understanding of the molecular mechanisms that control embryogenic

competence acquisition, somatic embryo development and conversion into viable plants. Nevertheless,

most of the results obtained are based on the traditional model systems, very often not easily compared

with the somatic embryogenesis systems of economical relevant woody species. The aim of this work is to

summarize some of the applications of proteomics in the understanding of particular aspects of the somatic

embryogenesis process in broad-leaf woody plants (model and non-model systems).

Key words Angiosperms , 2D electrophoresis , Embryogenic competence , Embryo maturation , Heat-

shock protein s , Mass spectrometry analysis , Metabolism-related protein s , Stress-related protein s ,

Zygotic embryo

1 Introduction

Proteomics studies the total proteins expressed in any given sys-

tem, whether by abundance, activity, structure, state of posttrans-

lational or other modiﬁ cation, or how these proteins interact with

each other in networks or complexes [

1 ]. In recent years proteome

studies have been employed to generate reference maps of the

most abundant soluble proteins of plant organs, at deﬁ ned devel-

opmental stages, for several purposes such as genetic studies com-

paring the proteomes of different plant genotypes, physiological

studies analyzing the inﬂ uences of exogenous signals on a particu-

lar plant organ, and developmental studies investigating proteome

changes during development [

2 , 3 ]. Technical advances provide

now a proteomic dissection of individual cell types, thus greatly

increasing the information revealed by proteome analyses [

2 ].

118

Proteomics has been successfully applied to the systematic anal-

ysis of protein expression during somatic embryo formation and

development in various plant species [

3 ]. Following the pioneering

studies in carrot [

4 , 5 ], somatic embryogenesis (SE) has been con-

sidered not only as an efﬁ cient system for in vitro clonal propaga-

tion, but also as an outstanding model system quite appropriate to

better understand totipotency in higher plants, as well as embryo

development , considering the difﬁ culties that have been encoun-

tered to analyze the early stages of zygotic embryo genesis during

development of the embryo inside the ovular tissues [

6 ].

Nevertheless, most knowledge on the general principles underlin-

ing the SE regulatory pathways has been focused on traditional

model organisms. With the recent increase in the number of

genome-sequencing projects, the deﬁ nition of model organism has

broadened [

7 ]. For example, the whole sequence genome of

Populus trichocarpa was published in 2006 [

8 ]. In addition, several

other tree species have been sequenced, including conifers [

9 – 11 ],

Eucalyptus [

12 ], and Fagaceae [ 13 ]. Moreover, the genomic data

for fruit trees such as citrus or apple also became available [

14 , 15 ].

Considering that there are approximately 300,000 botanically

described species of plants and that model plants represent only a

handful of species and families, even the arrival of these new model

plants cannot reﬂ ect the biodiversity of the plant kingdom and all

the economic or agricultural interests [

16 ]. Some features and pro-

cesses are unique and cannot be approached via a model plant.

Woody plants for example, are perennials with a quite long life cycle

and special features to be analyzed, including in what concerns their

SE systems. Proteomic approaches have a great potential to study

non-model species, because protein sequences have the advantage

of being more conserved, making the high- throughput identiﬁ ca-

tion of non-model gene products quite effective by comparison to

orthologous proteins [

17 ]. However, it is important to recognize

that there is a possible discrepancy between the messenger (tran-

script) and its ﬁ nal effector (mature protein). As most biological

functions in a cell are executed by proteins rather than by mRNA,

transcript expression proﬁ ling does not always provide pertinent

information for the description of a biological system. Several post-

transcriptional and posttranslational control mechanisms such as

the translation rate, the half-lives of mRNAs and proteins, protein

modiﬁ cations and intercellular protein trafﬁ cking, have an impor-

tant inﬂ uence on the phenotype [

18 ].

The main goal of this work is not to give a full review of all the

proteomic studies carried out on broad-leaf woody plant species,

but to summarize some of the applications of proteomics to under-

stand different aspects of the SE process in these plants (model and

non-model systems). First, a general perspective of the most com-

mon methodologies followed in the proteomics workﬂ ow is given,

followed by a short review of what we could learn in the last years

Sandra I. Correia et al.

119

from the proteomic approaches to embryogenic competence

induction and somatic embryo maturation and germination in

woody plants.

2 Proteomics Workﬂ ow

The most common proteomic workﬂ ow (Fig. 1 ) consists of pro-

tein extraction, protein (peptide) separation and quantiﬁ cation,

protein identiﬁ cation, and data integration [

19 , 20 ]. Several

approaches have been developed to address proteomic investiga-

tions, either trough top-down or bottom-up strategies, applying

“gel-based” or “gel-free” procedures. These procedures differ in

the way proteins are isolated (extracted), separated, and detected,

and consequently, each of them covers a typical subset of proteins

[

19 ]. The “gel-based” approaches based on two-dimensional poly-

acrylamide gel electrophoresis (2DE-PAGE) are the most com-

mon referred proteome analysis of the plant SE process. The

classical 2DE protocol separates denatured proteins according to

two independent properties: isoelectric point (p I ), by isoelectric

focusing (IEF), and molecular weight (MW). One of the most

challenging steps of the process is usually protein extraction from

plant samples, due to the relatively low protein content and high

level of contaminants [

21 ]. The cell wall and the vacuole are associ-

ated with numerous substances responsible for irreproducible

results such as proteolytic breakdown, streaking and charge het-

erogeneity. Most common interfering substances are phenolic

compounds, proteolytic and oxidative enzymes, terpenes, pig-

ments, organic acids, ionic species and carbohydrates . The majority

of the plant protocols introduce a precipitation step to concentrate

the proteins and to separate them from the interfering compounds.

The most commonly used method for extraction of plant proteins

is the trichloroacetic acid (TCA)/acetone precipitation method

[

22 ]. Apart from the optimization of the extraction protocol, also

protein solubilization is a critical factor. Proteins are solubilized in

the presence of high concentrations of chaotropes, a reductant and

a neutral detergent. The use of a detergent in conjunction with

chaotropes is of paramount importance and is decisive for the sub-

set of proteins that can be analyzed [

20 , 21 ]. Proteins of several

samples can be labeled prior to an electrophoretic separation with

spectrally distinct ﬂ uorescent dyes, and mixed together to run on

the same 2D gel. This 2D difference gel electrophoresis (DIGE)

approach allows to simultaneously comparing the proteomic pro-

ﬁ les of different samples that migrate under identical conditions

[

20 ]. After separation through 2DE, data are generated through

image analysis software that detects and quantiﬁ es the protein

abundances and matches the proteins ac ross the different gels.

Though the matching quality is dependent on the software

Proteomic Analysis of Somatic Embryogenesis in Broad-Leaf Woody Plants

120

Fig. 1 Workﬂ ow for the proteomic analysis of somatic embryo genesis in woody plants

Sandra I. Correia et al.

121

algorithm, it is above all determined by the quality and reproduc-

ibility of the gels. The standard approach for the identiﬁ cation of

2DE-separated proteins involves an enzymatic digestion of the

protein in the spot of interest and extraction of the peptides fol-

lowed by mass spectral (MS) analysis. The traditional way of analy-

sis involves peptide mass ﬁ ngerprint (PMF) analysis, typically

performed by matrix-assisted laser desorption/ionization-time of

ﬂ ight (MALDI-TOF) MS, since it provides a simple proﬁ le by pro-

ducing a single peak per peptide [

19 ]. However, PMF data have

very little resolution power to identify proteins from species with a

fragmentary genome and protein repository (non-model organ-

ism). Hence, the chance of ﬁ nding signiﬁ cant and conserved pep-

tides decreases and PMF fails or results in false positive hits [

23 ].

For that reason, tandem mass spectrometry (MS/MS) has been

often used to generate sequence speciﬁ c information and the infor-

mation content of such spectra is thus much higher than for

PMF. Unfortunately, separation of peptides prior to MS/MS is

expensive and time consuming, and MALDI-TOF is often pre-

ferred because of easiness of using, speed and ability to include

MALDI-TOF spotting in automated digestion protocols on liquid

handling systems [

19 ].

An emerging method gaining popularity combines one-

dimension (1D) gel separations with reversed-phase (RP) liquid

chromatography. Here proteins are ﬁ rst separated by size on stan-

dard polyacrylamide gels or by isoelectric point on IPG strips, nor-

mally used for the ﬁ rst dimensional separation in 2D-PAGE. After

separation, the lane of the gel or the strip containing the proteins

is extracted, divided into slices and treated similarly to spots excised

from 2D gels. The peptides are then separated on an integrated

and reusable RP column coupled to a standard HPLC pump. The

RP eluent is then analyzed by MS/MS [

24 ]. Although the plat-

form based on 2-DE is still the most commonly used [

25 ], the use

of “gel-free” approaches offers several advantages, since 2-DE is

difﬁ cult to automate. Most of the protocols use a bottom-up strat-

egy where proteins are ﬁ rst digested with a proteolytic enzyme and

the obtained complex peptide mixture is then separated via

reversed-phase (RP) chromatography coupled to a tandem mass

spectrometer [

24 ]. The whole dataset of acquired tandem mass

spectra is subsequently used to search protein databases and to link

the individual peptides to the original proteins. However, this con-

cept is only successful when identifying proteins in relatively simple

mixtures. In general, such peptide centered bottom-up approaches

have the disadvantage that both qualitative and quantitative infor-

mation on protein isoforms and differential posttranslational mod-

iﬁ cations are lost [

20 ].

To summarize, an optimized workﬂ ow for a non-model organ-

ism comprises (1) the investment in a powerful protein extraction

method capable to minimize the effects of interfering compounds,

Proteomic Analysis of Somatic Embryogenesis in Broad-Leaf Woody Plants

122

(2) the combination of different complementary protein fraction-

ation, separation and quantiﬁ cation techniques to maximize the

resolution and to cover the proteome as good as possible, and (3)

the usage of different complementary MS techniques and error

tolerant database searches [

19 ].

3 Proteomics Approaches to Somatic Embryogenesis Analysis in Broad-Leaf

Woody Plants

Proteomic studies have shown to be powerful tools for monitoring

the physiological status of plant organs under speciﬁ c developmen-

tal conditions [

3 ]. SE is one form of non- zygotic embryo genesis

by which somatic cells, under suitable induction conditions,

undergo a complete genome shift and embark into a new develop-

mental pathway ending in the formation of asexual embryos mor-

phologically identical to their zygotic counterparts [

6 , 26 , 27 ].

During this unique developmental process, cells have to dediffer-

entiate, activate cell division, and reprogram their physiology,

metabolism and gene expression patterns [

28 ]. Thus, SE can be

considered the clearest demonstration of totipotency , showing that

somatic cells contain the essential genetic blueprint to complete

plant development, and that embryogenesis is not exclusive of the

zygote formation and can proceed in absence of fertilization [

26 ].

Since the ﬁ rst observations of somatic embryo formation in car-

rot cell suspension cultures [

4 , 5 ], the potential for SE has been

shown to be characteristic of a wide range of tissue culture systems

from both gymnosperms and angiosperms plants [

28 – 30 ].

In recent years, there has been a growing interest in proteomic

approaches to better understand SE. Since proteins directly inﬂ u-

ence cellular biochemistry and provide a more accurate analysis of

cellular changes during growth and development [

31 ], the identi-

ﬁ cation of proteins associated with somatic embryo development

may provide insights onto SE. Thus, several proteomic approaches

were applied to study somatic embryogenesis of several broad-leaf

woody plant species such as cork oak ( Quercus suber ) [

32 ], Valencia

sweet orange ( Citrus sinensis ) [

33 ], grape wine ( Vitis vinifera )

[

34 ], cacao tree ( Theobroma cacao ) [ 35 ], feijoa ( Acca sellowiana )

[

36 ], and tamarillo ( Cyphomandra betacea ) [ 37 ]. These reports

included studies on protein expression changes during SE and

comparative studies between embryogenic and non-embryogenic

cells as well as between zygotic and somatic embryogenesis.

SE induction involves differentiated somatic cells acquiring

embryogenic competence and proliferating as embryogenic cells

[

28 ]. This switching of somatic cells into embryogenic cells

involves a series of events associated with the molecular recogni-

tion of internal signals and external stimuli [

38 , 39 ]. In recent

3.1 Embryogenic

Competence

Acquisition

Sandra I. Correia et al.

123

years, an increasing number of works have indicated that the stress-

response of cultured tissues plays a major role in somatic embryo

induction [

39 , 40 ], and that plants respond to abiotic stresses by

altering the expression of many of their genes. This altered expres-

sion is a major mechanism of adaptation and survival during the

stress periods [

41 ]. Actually, proteomics helps the investigation of

changes in proteome proﬁ les emphasizing the participation of

stress-related protein s in all developmental processes [

3 ].

One important line of investigation to analyze embryogenic

competence acquisition by woody plants is by the comparison of

responsive and nonresponsive explants during the SE induction

process [

37 , 42 – 44 ]. In the late 1990s, following the pioneering

works of De Vries and collaborators in carrot [

45 ], the detection

of embryogenesis-related proteins from total protein extracts was

reported for several woody species, such as Betula pendula [

41 ],

Camelia japonica [

46 ], and Cupressus sempervirens [ 47 ]. In Betula

pendula , the changes in protein patterns and the expression of

“embryo-speciﬁ c” proteins during embryogenesis were observed

when comparing two cell lines, one potentially embryogenic,

under the right inductive conditions, and one which never has

shown any embryogenic capacity. In the following years, the

improvements in high-resolution 2-DE and mass spectrometry

contributed to the large-scale proﬁ ling and identiﬁ cation of the

proteins associated to embryogenic competence acquisition. SE

systems in which embryogenic (EC) and non-embryogenic (NEC)

cell lines can be induced from the same cultured explant, like the

ones of wine grape ( Vitis vinifera ) [

34 , 43 ] and tamarillo

( Cyphomandra betacea ) [

37 ], have been explored to obtain more

information on important regulatory proteins. Proteins, exclu-

sively or predominantly expressed in EC, included iron-deﬁ ciency-

responsive proteins, acidic ascorbate peroxidases and isoﬂ avone

reductase-like proteins [

43 ] and metabolism-related protein s , such

as enolases and threonine synthases, and also heat-shock protein s

(HSP) and ribosomal proteins [

37 ]. Ascorbate peroxidases, cata-

lases, calcineurin B-like proteins, 1,3- b -glucanases, cyclin-

dependent kinases A1 [

43 ] and pathogenesis-related (PR) proteins

were found mainly in NEC [

37 , 43 ]. The examination of differen-

tially expressed proteins between ECs and NECs suggests that the

embryogenic status of EC cells could be related to a better ability

to regulate the effects of stress conditions, namely through the

controlling of oxidative stress by regulation of the reactive oxygen

species ( ROS ) scavenging system [

34 ], and by the action of HSP

[

34 , 37 ]. A hypothesis is that the expression of totipotency in cul-

tured somatic cells is part of a general stress adaptative process that

implies a ﬁ ne regulation of auxin and stress signaling resulting in

the restart of cell division and embryogenic competence acquisi-

tion. The observation that embryogenic tissues of different origins

and obtained with the use of different auxins display similar pro-

tein proﬁ les suggests a general behavior of cellular metabolism that

Proteomic Analysis of Somatic Embryogenesis in Broad-Leaf Woody Plants

124

can give important insights about the mechanisms triggering and

controlling somatic embryo formation [

37 ]. Also for cork oak

[

32 ], the role of ROS in the proliferative stages during SE and the

upregulation of proteins involved in cell division were reported.

The comparison between somatic embryo cells type (SE-type) and

pro-embryogenic masses type ( PEM -type) of avocado ( Persea

americana ) have conﬁ rmed the observations previously made in

other systems [

44 ]. In this work, the identiﬁ cation of high levels of

HSP, glutathione S -transferases (GST), and superoxide dismutases

(SOD) proteins in SE-type cells suggested that the generation of a

signiﬁ cant amount of stress and ROS are prerequisites to induce

somatic embryogenesis, and SE lines seems to be more efﬁ cient to

cope with the necessary ROS and stress and, hence, have a higher

regeneration capacity.

In order to develop into somatic embryo s, somatic cells must

regain their cell division activity. Hence, the division associated

proteins, such as proliferating cell nuclear antigen in grape wine

[

34 ] and putative citrus DRT102 in Valencia sweet orange [ 33 ]

are activated during embryogenesis. Besides, cytoskeletal proteins,

such as tubulins associated to cell division, are also differentially

regulated [

33 ]. During the last decades, proteomic studies also

described several extracellular proteins as markers for SE, which

could offer the possibility of determining embryogenic potential of

plant cells in culture [

38 – 49 ]. Arabinogalactan protein s , nonspe-

ciﬁ c lipid transfer proteins and germin/germin-like proteins are

important groups of extracellular proteins that help triggering

embryogenic potential in plant cells [

50 ]. More recently, results

obtained with EC and NEC suspension cultures of coffee species

( Coffea sp.) [

51 ] showed that a particular set of proteins is exclu-

sively secreted under embryogenic conditions.

In several plant regeneration processes through SE, one of the

major problems is an effective transition from the proembryogenic

masses, forming the embryogenic tissue, toward embryo develop-

ment , which is often impaired by the formation of abnormal

embryos and precocious germination of many others. This situa-

tion may be caused by an inadequate maturation of the embryos,

an important phase of somatic and zygotic embryo development,

following the classic morphogenic phases from globular to

cotyledonary embryos [

52 ]. During maturation, embryo cells

undergo various physiological changes, which become evident by

the deposition of storage materials, repression of germination and

acquisition of desiccation tolerance [

53 , 54 ]. In cork oak, the acti-

vation of diverse ROS detoxiﬁ cation enzymes and the accumula-

tion of reserve products ( carbohydrates and proteins mostly) have

been reported during the transition phase between proliferation

and cotyledonar stages, suggesting the requirement that cell divi-

sion should be replaced with cell expansion for proper embryo dif-

ferentiation [

32 ]. Energy requirements reach a maximum at the

3.2 Somatic Embryo

Maturation

and Conversion

Sandra I. Correia et al.

125

cotyledonary stage, suggesting the relevance of primary metabolite

production, such as amino acids and fatty acids, whereas fermenta-

tion could constitute an alternative source of energy at the early

steps of somatic embryo development [

32 ]. Also, for Valencia

sweet orange [

33 ] several proteins involved in antioxidative stress

response (GST), cell division (tubulins), photosynthesis (ferritins),

and cyanide detoxiﬁ cation (rhodanese) exhibited different expres-

sion patterns and were likely to be associated with SE. Another

species often referred in studies aimed to detect and identify pro-

teins expressed during the different developmental stages of

somatic embryos is the myrtaceous feijoa ( Acca sellowiana ) [

36 ,

55 ]. The results obtained with this SE system indicate a high simi-

larity in the proﬁ les of the assayed somatic embryos, suggesting

that only a few speciﬁ c genes are involved in the different develop-

mental stages, and that gene expression occurs prior to morpho-

logical changes. The hypothetical protein similar to

l -isoaspartyl- O -methyltransferase in torpedo stage, and an osmotin-

like protein in the pre-cotyledonar stage of somatic embryos were

suggested as embryonic markers for feijoa [

55 ]. The expression of

the protein phenylalanine ammonia lyase in all the assayed devel-

opmental stages conﬁ rmed the synthesis and accumulation of sev-

eral phenolic compounds observed during the induction of feijoa

embryogenic cultures and the development of somatic embryos.

The presence of cytosolic glutamine synthetase and NmrA-like

proteins revealed the activation of nitrogen metabolism, observed

particularly in the later developmental stages in which the accumu-

lation of storage compounds (mostly in the cotyledonary leaves) is

enhanced [

55 ]. More recently, the comparison between “off-type”

and normal phenotype proteomes of somatic plantlets of feijoa has

brought new insights to somatic embryo abnormal development

[

36 ]. The presence of HSP was observed only during the forma-

tion of normal phenotype somatic plantlets, indicating that these

proteins may be involved in the morphogenesis of normally devel-

oped plantlets. A vicilin-like storage protein was only found in

“off-types” at 20-day conversion , indicating that plantlets may

present an abnormality in the mobilization of storage compounds,

causing reduced vigor in the development of derived plantlets.

The understanding of seed development is an important approach

to overcome the difﬁ culties in somatic embryo conversion and ger-

mination . Proteomic analyses have been made on zygotic embryo s

of several woody species, such as araucaria ( Araucaria angustifolia )

[

56 ], coffee ( Coffea arabica ) [ 57 ], and Masson’s pine ( Pinus mas-

soniana ) [

58 ]. These analyses revealed signiﬁ cant requirements for

energy production/carbon metabolism during the early stages of

zygotic embryo development [

35 , 59 ]. Over accumulated proteins

in early seed development also indicated a higher control on oxida-

tive stress metabolism during this phase [

56 ]. Besides, early zygotic

embryos showed changes in the abundance of proteins involved in

3.3 Somatic Versus

Zygotic Embryos

Proteomic Analysis of Somatic Embryogenesis in Broad-Leaf Woody Plants

126

mRNA splicing and signaling [ 57 ]. The advanced stages of zygotic

embryogenesis were complemented by differential expression of

Rubisco, Myb transcription factor , and by changed biosynthetic

activity of phosphatidylcholine [

57 ], but also by an active metabo-

lism, leading to carbon assimilation and storage compounds accu-

mulation [

56 ]. Comparison of somatic and zygotic embryos

revealed that their proteomes reﬂ ected mainly the different envi-

ronmental conditions, which caused differential expression of pro-

teins involved in metabolic pathways and stress response [

59 ].

Noah and collaborators systematically compared at the proteome

level the physiological mechanisms underlying somatic and zygotic

embryogenesis in cacao [

35 ]. Many of the identiﬁ ed proteins were

involved in genetic information processing, carbohydrate metabo-

lism and stress response. Somatic embryo s especially displayed

many stress related proteins, few enzymes involved in storage com-

pound synthesis and an exceptional high abundance of endopepti-

dase inhibitors. Phosphoenolpyruvate carboxylase, which was

accumulated more than threefold higher in zygotic embryos, rep-

resents a prominent enzyme in the storage compound metabolism

in cacao seeds. More information on this topic are reported else-

where in this book ( see Chapter

2 ).

4 Conclusions and Future Perspectives

The results obtained with the proteomic studies over the last

decades strongly emphasize the role of stress proteins in somatic

embryo genesis, revealing an intricate dynamism, variability, and

behavior of several regulatory proteins. Nevertheless, and unlike

the classical biological model systems, the full potential of pro-

teomics is far from being fully exploited in woody plant research.

Only a low number of woody species has been investigated at the

proteomics level and the predominant use of strategies based on

2DE coupled to MS results have so far resulted in a low proteome

coverage. Although proteome analyses are still signiﬁ cantly less

representative in the literature than those based on genomic

approaches, the integration of the expressed protein data, together

with transcriptome and even metabolome data, has the potential to

provide the most comprehensive and informative clues on somatic

embryogenesis in plants. Future research in this ﬁ eld should

include new and/or complementary approaches, including more

sensitive methods for protein detection and identiﬁ cation. The

laser-capture microdissection tool is one of those technical improve-

ments that could overcome the limitations in tissue accessibility,

allowing more accurate molecular proﬁ les of isolated embryogenic

tissues. Also, “gel-free” approaches, with higher levels of automa-

tion and less inherent technical variability, should be applied in

order to obtain a better reproducibility. These approaches are

becoming more effective with the integration of new data from

Sandra I. Correia et al.

127

several genome-sequencing projects. Furthermore, efforts should

be taken in the functional validation of the speciﬁ c identiﬁ ed pro-

teins, in order to use them as markers for the SE process. The

coordination of all this knowledge will give an insight into future

studies addressing the optimization of the somatic embryogenesis

protocols for mass propagation and conservation strategies in sev-

eral economical relevant woody species.

Acknowledgements

This work was supported by a postdoctoral research fellowship

(SFRH/BPD/91461/2012) awarded to Sandra Correia by the

Fundação para a Ciência e Tecnologia (Portugal).

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Proteomic Analysis of Somatic Embryogenesis in Broad-Leaf Woody Plants

131

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_7, © Springer Science+Business Media New York 2016

Chapter 7

Advances in Conifer Somatic Embryogenesis

Since Year 2000

Krystyna Klimaszewska , Catherine Hargreaves , Marie-Anne Lelu-Walter ,

and Jean-François Trontin

Abstract

This review compiles research results published over the last 14 years on conifer somatic embryogenesis

(SE). Emphasis is placed on the newest ﬁ ndings that affect the response of seed embryos (typical explants)

and shoot primordia (rare explants) to the induction of SE and long-term culture of early somatic embryos.

Much research in recent years has focused on maturation of somatic embryos, with respect to both yield

and quality, as an important stage for the production of a large number of vigorous somatic seedlings.

Attempts to scale up somatic embryo production numbers and handling have resulted in a few bioreactor

designs, the utility of which may prove beneﬁ cial for an industrial application. A few simpliﬁ ed cryopreser-

vation methods for embryonal masses (EM) were developed as a means to ensure cost-efﬁ cient long-term

storage of genotypes during clonal ﬁ eld testing. Finally, recent long-term studies on the growth of somatic

trees in the ﬁ eld, including seed production yield and comparison of seed parameters produced by somatic

versus seed-derived trees, are described.

Key words Cryopreservation , Field tests , Somatic embryo s , Somatic trees , Tree improvement

1 Discovery of Somatic Embryogenesis in Conifers and the General Pattern

of Somatic Embryo Development

Since its discovery in Picea abies and Larix decidua [ 1 , 2 ], somatic

embryo genesis (SE) in conifers has been reported in many other

species, with a large majority of them belonging to Pinaceae,

and only a few to Cupressaceae , Taxaceae , Cephalotaxaceae , and

Araucariaceae families. The general differentiation and develop-

mental pattern of conifer somatic embryos is highly similar among

most species tested to date and starts with an immature seed

embryo (enclosed in a megagametophyte or excised) or with an

excised mature embryo that is cultured on a nutrient medium con-

taining plant growth regulator s (PGRs) from either both the auxin

and cytokinin groups or from cytokinin only. Subsequently, the

132

cells of the embryo deviate from their previous pattern of division

and differentiation into a mature embryo; instead, they start divid-

ing profusely and differentiate into multiple early somatic embryos

also known as embryonal masses ( EM ). For example, 50 mg fresh

mass of white spruce proliferating cell culture may contain, at any

given time, approximately 300 single early somatic embryos, 39–75

cleaving early somatic embryos, 60–90 multiple cleaving somatic

embryos, small and large cell aggregates, and single small and large

cells, the latter with large vacuole(s) (Klimaszewska, unpublished).

Typically, the composition of the culture is highly heterogeneous

and may change over culture time, a period that may last from

several months to several years, during which EM has to be subcul-

tured every 10–21 days (depending on the species) onto fresh

medium of the same or slightly modiﬁ ed composition. EM of some

species can be cultured on a semisolid medium or in liquid medium.

The cultures are amenable to long-term storage through cryo-

preservation without losing their viability and growth characteris-

tics. In most species, early somatic embryos will not develop further

unless the culture conditions are changed. High frequency devel-

opment and maturation of early somatic embryos in most conifer

species of the Pinaceae family take place under remarkably similar

conditions, namely in the presence of abscisic acid ( ABA ), sucrose

and/or other sugars, and in a medium that imposes restriction on

water availability either by physical means (high concentration of

solidifying agents) or by a high molecular weight solute such as

polyethylene glycol ( PEG , MW 4000–8000) that mimics drought.

Often the sugar concentration is also increased to lower the osmotic

water potential of the medium. Once the somatic embryos resem-

ble mature seed embryos (usually after 6–12 weeks, depending on

the species), they are harvested and desiccated if matured on a

medium with PEG for normal germination , or they can be germi-

nated directly if developed on a medium with high gel strength.

On a germination medium, the somatic embryos display a rapid

radicle and hypocotyl elongation (usually within 1 or 2 days) fol-

lowed by the growth of a shoot and a root. Once the plantlets

reach the desired size, they are transferred into containers with a

suitable substrate and acclimatized in a greenhouse or a nursery.

Subsequently, they are planted in the ﬁ eld for research purposes,

clonal selection or, eventually, for commercial production.

Several reviews describing various aspects of SE research in

conifers have been published in both international journals and in

books [

3 – 9 ]. The main focus of the present review is to summarize

the progress in SE research in conifers made over the last 14 years

and to determine its impact on understanding the basic mecha-

nisms governing the process and on the development of new, more

efﬁ cient protocols for the production of somatic trees. We also

include research results on conifer species for which somatic

embryo genesis has been achieved only recently and the results

obtained with adult trees.

Krystyna Klimaszewska et al.

133

2 Genetic Control in Somatic Embryogenesis

Perhaps the most basic factor that determines whether SE is

initiated from the seed embryos is the genetics of parental trees,

providing that suitable culture medium and conditions have been

established. Experiments designed to establish the extent of genetic

control in SE initiation have been conducted with a few pine spe-

cies. A large study was undertaken with Pinus sylvestris using 49

seed families from diallel crosses among seven elite trees including

reciprocals and selﬁ ng [

10 ]. Four of the experimental trees were

preselected for their propensity for SE based on an earlier study

that tested 138 trees. Analysis of the data suggested a stronger

maternal than paternal effect on culture initiation; however, spe-

ciﬁ c combining ability (SCA) had no detectable effect. The mater-

nal effect at the initiation stage could be explained by both the

genotype and the developmental or physiological stage of the

mother tree and the inherited maternal alleles of the zygotic

embryo . Similarly, MacKay et al. [

11 ] quantiﬁ ed the genetic con-

trol of SE initiation in P. taeda using seeds from diallel crosses and

factorial matings. Thirty seed families were used in the experiments

that tested two different culture treatments and resulted in large

differences between treatments in SE initiation frequency among

families. The variance due to treatments accounted for 41 % of

total phenotypic variance, whereas that due to families accounted

for 22 %. Signiﬁ cant variance due to interactions between families

and treatments was also found, accounting for 13 % of the pheno-

typic variance. The latter indicated that different culture media

might be better suited for different genotypes. In another study

with 20 control-pollinated seed families of P. radiata , Hargreaves

et al. [

12 ], also challenged the notion that poor results should be

attributed to genetic effects only and showed that it was possible to

create laboratory conditions that increased the number of respond-

ing explants ac ross all families. However, previous work with P.

taeda showed that many mother trees produce seeds that do not

respond to SE initiation, leading to the hypothesis that such trees

possess unfavorable alleles at loci expressed in the mother tree,

whereas favorable alleles at other loci may be inherited by zygotic

embryos [

11 ]. The estimates of large general combining ability

(GCA) variance component and narrow sense heritability sug-

gested that targeted breeding could inﬂ uence SE initiation in P.

taeda . To test this hypothesis, an experiment was carried out that

involved a small number of control reciprocal crosses among trees

that ranged from low to high SE initiation capacity when tested

with seed from open-pollinated mother trees. By selecting a favor-

able mother tree for cross-pollination for each pair of parental

trees, it was possible to increase SE initiation frequency from 1.5-

to 9-fold. Also, some trees had strong additive effects as male parents,

but had negative maternal effects; hence, using them in control

Conifer Somatic Embryogenesis

134

crosses as pollen donors might be yet another solution to increase

SE initiation. The authors concluded that this knowledge of

genetic control in SE initiation can now be easily applied to breed-

ing schemes to capture valued genotypes. Smaller studies with

seeds from control crosses of P. pinaster [

13 ] and P. sylvestris [ 14 ]

supported the above conclusions.

3 Improvements of Previously Established Protocols for Somatic Embryogenesis

SE biotechnology of conifers has constantly evolved since its dis-

covery in 1985, and incremental improvements are being made

according to the time and effort committed to a given species. The

literature search revealed that since the year 2000, about 46 jour-

nal articles reporting improvements of SE protocols were pub-

lished for Pinus taeda , Pinus strobus , Pinus sylvestris , Pinus pinaster ,

Pinus radiata , Pinus patula , Pseudotsuga menziesii , Abies nordma-

nniana , Picea abies , Picea glauca , Picea mariana , and Larix

hybrids. In addition, the publication of approximately 40 articles

on species for which SE is described for the ﬁ rst time is a clear indi-

cator of the importance of this technology for conifer clonal propa-

gation. For species of economic importance that are grown in

forest plantations, a lot of research has been carried out by compa-

nies and patented, for example for Pinus taeda (Arbogen,

Weyerhauser, WA, USA), P. radiata (Forest Genetics Ltd and

Arborgen, NZ), P. abies , P. pinaster , and P. radiata (FCBA,

France), Pseudotsuga menziesii (Weyerhauser, WA, USA), Picea

glauca and P. abies (Natural Resources Quebec, QC, Canada and

JD Irving Inc., NB, Canada). Among these economically valued

plantation species, the largest body of literature exists for Pinus

taeda (loblolly pine), which reports on the stepwise optimization

approach to overcome low efﬁ ciencies at each stage of SE. In their

recent review, Pullmann and Bucalo [

9 ] have attributed these

improvements to medium supplements including speciﬁ c sugars,

vitamins, organic acids, and redox potential modiﬁ ers. Other con-

trolled factors, including medium water potential, pH, adsorption

of medium components by activated carbon and use of liquid ver-

sus semisolid medium, also positively inﬂ uenced SE. These modiﬁ -

cations resulted from the analytical studies of P. taeda seed tissue,

the seed environment, and gene expression in megagametophytes,

zygotic embryo s, and somatic embryo s. The premise of the study

was that duplication of the seed environment in vitro would lead to

the design of efﬁ cient protocols for SE.

Major improvements were made in the frequency of SE initiation

in either open-pollinated or control-pollinated seed sources of sev-

eral European and North American Pinus species, and Pseudotsuga

menziesii . In P. strobus , the number of responding immature seed

3.1 Initiation of SE

and Growth of Early

Somatic Embryos

Krystyna Klimaszewska et al.

135

embryos increased from the average of 20 % to the average of 53 %

ac ross ﬁ ve open-pollinated seed families by reducing the

2,4- dichlorophenoxyacetic acid (2,4-D) and benzyladenine (BA)

concentrations from 9.5 to 2.2 μM and from 4.5 to 2.2 μM,

respectively. Both concentrations were tested in modiﬁ ed [

15 , 16 ]

Litvay’s medium (MLV). The most striking difference in initia-

tions occurred when the embryos were at the pre-cleavage and

early post-cleavage stages, which were also linked to the morpho-

logical appearance of the megagametophytes becoming opaque as

opposed to being translucent [

15 ]. The same medium modiﬁ ca-

tions were tested with a few control-pollinated seed families of P.

sylvestris and the result was better on a medium with reduced

PGRs, i.e., 24 % initiation versus 9 % [

14 ]. However, contrary to

the response of P. strobus and P. sylvestris , when eight control-

pollinated seed families of P. pinaster were tested, MLV with

reduced concentrations of PGRs decreased the initiation frequency

from 93 to 80 %; nevertheless both culture medium variants were

very productive [

13 ]. The high response of the latter was also

attributed to the selection of embryos that were at the uniform

pre-cotyledonary stage of development by excising the embryos

from the surrounding megagametophytes. When the embryos

were cultured within megagametophytes, the SE response was

only slightly reduced, suggesting that in the tested cones the devel-

opment of embryos was relatively synchronized. This was in con-

trast to P. radiata for which the zygotic embryo s had to be excised

for the best response [

17 ].

Another medium that is considered suitable for SE initiation

in P. pinaster and P. sylvestris is Gupta and Durzan’s medium

(DCR [

18 ]). Like other commonly used media [ 7 , 19 , 20 ], it also

includes glutamine and casein hydrolysate as well as 2,4-D and

BA. Recent research aimed at improving SE initiation in P. radi-

ata showed that by making another modiﬁ cation to Litvay’s

medium (designated as GLITZ) it was possible to achieve consid-

erably higher responses from both 19 open-pollinated and 20

control-pollinated seed families compared with those obtained on

Embryo Development Medium 6 (EDM6) [

12 , 17 , 21 ]. Average

initiations were 70 % for both types of seed families when embryos

were excised from the megagametophytes at an early stage of

development. GLITZ medium contained glutamine (0.5 g/L),

casein hydrolysate (1.0 g/L), 2,4-D (4.5 μM), and BA (2.25 μM).

Likewise, a 2-year study on SE initiation in P. nigra demonstrated

higher potential for two out of four tested medium formulations

[

22 ]. DCR and MLV media consistently supported approximately

10 % explants producing EM as opposed to Litvay medium (LV

[

16 ]) and Quoirin and Lepoivre (QP [ 23 ]) media, on which the

response was negligible.

Among pine species, SE in P. taeda has been the most

researched owing to its high commercial value in the USA and

Conifer Somatic Embryogenesis

136

elsewhere, but relatively low responses obtained in earlier work.

Not surprisingly, a considerable effort was undertaken to improve

the efﬁ ciencies of SE and understand the bottlenecks at each stage

( see ref.

9 and references therein). Various supplements were tested

in a unique loblolly pine (LP) medium formulation and found ben-

eﬁ cial for SE initiation, such as AgNO

3, maltose instead of sucrose ,

high level of myo-inositol (up to 20 g/L), glutamine , casamino

acids, 2-( N -morpholino) ethanesulfonic acid (MES, as pH stabi-

lizer), biotin, folic acid, vitamins B

12 and E, α-ketoglutaric acid,

kinetin together with BA, activated charcoal , abscisic acid ( ABA ),

brassinolide as well as

D -xylose and D - chiro -inositol [ 24 ]. The num-

ber of initiated SE cultures increased further by adding liquid over-

lays 14 days after placement of explants on gelled medium. This

technique allows replenishment or addition of nutrients and PGRs,

or adjustment of pH without disturbing the tissue.

All the medium supplements that were beneﬁ cial to P. taeda

were also tested with immature seed embryos of P. menziesii , which

differ from those of pine species by the lack of cleavage polyem-

bryony and the need for an embryo to be cultured while partially

excised and still attached to the megagametophyte by a suspensor

[

25 ]. These tests resulted in the development of an effective

medium formulation for initiation of SE in P. menziesii that

included activated charcoal , ABA , biotin, brassinolide, folic acid,

MES, pyruvic acid,

D -xylose, and D - chiro -inositol in addition to 1

g/L myoinositol, 0.5 g/L casamino acids, 0.45 g/L glutamine

and 2,4-D, BA, and kinetin [

24 , 25 ]. Tests with seeds from high-

value crosses conducted over 2 years gave initiation frequencies of

40 and 57 %, respectively. Based on the above results, a new

medium was designed for the culture of mature embryos of P. abies

that resulted in doubling SE initiation from around 14–30 % when

the medium contained 100 mg/L

D -xylose. Some other medium

additives were asparagine and brassinolide, and the PGRs were

α-naphthaleneacetic acid (NAA) and BA [

24 ].

Research on initiation of SE in somatic embryo s of Larix x

leptoeuropaea showed that 98 % of cotyledonary somatic embryos

matured for 3 weeks produced SE; those matured for 6 weeks pro-

duced SE at a frequency of only 2 % [

26 ]. The authors suggested

that the loss of ability of somatic embryos to respond to the

induction treatment might be caused by the synthesis/accumula-

tion of ethylene , because enrichment of the vessel headspace with

ethylene reduced the induction of SE from 3-week-old somatic

embryos from 98 to 4 %. Ethylene was also found to inﬂ uence the

development of early somatic embryos as described below.

Once SE is initiated and EM can be identiﬁ ed (after several days to

several weeks), the next challenge is to ensure a rapid proliferation

of EM upon subculture onto fresh medium to generate the

amounts that are needed for various steps, such as cryopreservation

3.2 Growth

of Initiated EM

Krystyna Klimaszewska et al.

137

and/or production of mature somatic embryo s. The majority of

conifer species are usually subcultured onto media of the same

composition, but in a few cases, such as P. pinaster , it has been

shown that medium modiﬁ cations were required to obtain better

growth and/or to maintain the embryogenic potential of the cul-

tures [

27 , 28 ]. These modiﬁ cations included weekly subcultures,

substitution of sucrose with maltose and withdrawal of PGRs

(2,4-D and BA). To maintain satisfactory growth of P. radiata

EM, it was necessary to increase the amino acid content in LV

medium after initiation [

12 ]. In some species and genotypes, the

application of a culture technique that is based on dispersing the

cells in a liquid medium, and then collecting the cells on a ﬁ lter

paper, draining the liquid, and placing the ﬁ lter paper with the cells

onto a fresh medium has been the most important for the survival

and growth of P. monticola and P. sylvestris [

14 , 29 ].

Embryogenic cultures of Cryptomeria japonica ( Cupressaceae )

were composed of a mixture of EM and callus cells, and when an

attempt was made to culture the EM separately; their embryo-

genic capacity was lost [

30 ]. The culture medium was that of

Campbell and Durzan [

31 ], containing 1 μM 2,4-D and 0.6 g/L

glutamine ; however, when the medium was supplied with 2.46

g/L glutamine, the culture remained embryogenic and simulta-

neously its dry mass and endogenous level of glutamine increased.

The high glutamine treatment might have increased the synthesis

of certain macromolecules or metabolites that were essential for

SE. The ability of EM to grow in the presence of callus cells was

attributed to the high content of endogenous glutamine in the

latter that might have supported the growth of EM in mixed cul-

ture. Based on the research results of others, the authors con-

cluded that without an adequate supply of glutamine/glutamate,

the embryogenic culture of C. japonica would lose its embryo-

genic characteristics. Phytosulfokine, which is a small sulfated

peptide, was also found beneﬁ cial for C. japonica culture growth

when included in the medium at 32 nM [

32 ]. This peptide acts as

an extracellular ligand at the onset of cell dedifferentiation, prolif-

eration, and redifferentiation and plays a stimulatory role in SE. In

particular, phytosulfokine promoted suspensor regeneration from

basal cells of somatic embryo s of Larix leptolepis [

33 ]. The pres-

ence of suspensor in somatic embryo development and in the

maintenance of the culture embryogenic characteristics was estab-

lished as critical by Umehara et al. [

34 ] and later by Larsson et al.

[

35 ] and Abrahamsson et al. [ 36 ] for Picea abies and Pinus sylves-

tris , respectively.

To promote further development and maturation of early somatic

embryo s in a majority of conifer species, the proliferating cultures

of early somatic embryos are transferred (after a pretreatment or

without it) onto a medium with ABA that replaces auxin and/or

cytokinin. Most often, the medium water potential is lowered at the

3.3 Development

and Maturation

of Early Somatic

Embryos

Conifer Somatic Embryogenesis

138

same time by increasing the concentration of sugar(s) and creating

permeating osmotic stress, or by including PEG (MW 4000–8000),

thus creating a non-permeating osmotic stress, the latter is due to

the larger than cellular pores molecule size [

37 ]. An alternative

method of affecting somatic embryo development is to increase the

gelling agent concentration in the ABA medium, which increases

gel strength and reduces water availability to the cells [

38 ].

A study that unequivocally conﬁ rmed that ABA was crucial for the

normal somatic embryo development and maturation in conifers

was carried out with Larix x leptoeuropaea [

39 ]. This larch hybrid

is somewhat unique because it can produce cotyledonary somatic

embryos and plantlets on a medium with ABA or without it, hence

providing an ideal material for this study. However, the somatic

embryos that developed on both media differed in structure, cell

types, intracellular secondary metabolites and storage product

accumulation, endogenous ABA concentrations, and extracellular

mucilage build-up. Clearly, those from ABA medium displayed a

coordinated growth and better-shaped somatic embryos with the

concomitant accumulation of lipids and storage proteins that were

lacking in embryos developed in the absence of ABA. Hence,

somatic embryos developed without ABA did not go through mat-

uration. Still, in all conifer species studied to date, a certain num-

ber of genotypes in a given species fails to produce mature somatic

embryos even in the presence of optimized concentrations of

ABA. It has been shown that the ability of embryogenic tissue to

utilize ABA from the medium may reﬂ ect the capability of embryo

maturation in different genotypes of Picea glauca x engelmanni

[

40 ]. The genotypes that produced mature somatic embryos on

gelled medium with racemic ABA (equal amounts of (+)- cis , trans-

ABA and (−)- cis , trans -ABA) were characterized by a greater utili-

zation of exogenous ABA, when grown as cell suspension s,

compared with a non-productive genotype. Furthermore, differ-

ent forms of ABA were metabolized to various levels. For example,

only half of racemic ABA was metabolized by the 22nd day of

culture; the remainder was exclusively (−)-ABA. The natural ABA

((+)-cis, trans-ABA) was still available at the end of the test, but its

amount may be inﬂ uenced by species, cell density of the initial

inoculation, tissue growth rates, and initial ABA concentrations.

The natural ABA exerted by far the best bio-effect compared with

racemic ABA and the mixture of ABA isomers.

Improvement of somatic embryo quality and yield was achieved by

combining ABA with activated charcoal ( AC ). In P. abies , AC

introduced into a medium at 0.125 % with 189 μM ABA promoted

a zygotic-like appearance of somatic embryos with more elonga-

tion and taper in the hypocotyl region as well as formation of a

prominent shoot apical region compared with those developed

3.3.1 Abscisic Acid

3.3.2 Activated Charcoal

Krystyna Klimaszewska et al.

139

without AC [ 41 ]. These embryos grew faster and were produced

at a reduced material and labor cost because the cultures did not

require subculturing onto fresh medium. However, the authors

cautioned against the types of AC to be utilized as these vary with

respect to particle sizes and hence the adsorption properties caus-

ing potential deﬁ ciencies in the medium components. Alternatively,

AC was used with P. pinaster by coating the cells with AC and

culture on a ﬁ lter paper placed on the maturation medium [

13 ].

This method of culture resulted in a greater number of mature

somatic embryos produced in a shorter time compared with cul-

tures without AC coating. Similarly, P. sylvestris aged cultures (24

weeks old) responded favorably when coated with AC, whereas

there was no effect on young cultures (8 weeks old) [

14 ].

Changes in water status that occur during conifer zygotic embryo

development and maturation are also critical for the progression of

the development of early somatic embryo s, but the type of com-

pounds used to alter the medium water status must be the “right”

type for a given species. For example, when mannitol (a plasmolyz-

ing agent) was tested against PEG (a non-plasmolyzing agent) in

cultures of P. glauca , the better quality of somatic embryos from

the latter was accompanied by the accumulation of higher levels of

reduced ascorbate, resulting in a physiological state similar to that

of zygotic embryos [

42 ]. Moreover, in the presence of PEG, there

was a constant decline in the GSH (reduced)/GSSG (oxidized)

ratio of glutathione, suggesting seed-like ﬂ uctuations of the

ascorbate- glutathione metabolism in somatic embryos. In another

study with somatic embryos of P. glauca and P. mariana it was

found that sucrose at 6 % (in the absence of PEG) was highly ben-

eﬁ cial when added to the maturation medium for both the number

of matured somatic embryos and the accumulation of soluble and

insoluble storage proteins [

43 ]. The maturation response could

not be matched by osmotic equivalents of glucose and fructose

(products of sucrose hydrolysis) in the medium. Moreover, the

embryo carbohydrate content was independent from the carbohy-

drate used in the maturation medium. The same conclusion was

later reached for somatic embryos of P. abies , where endogenous

carbohydrate patterns were stable irrespective of culture condi-

tions, which indicated the carbohydrate status to be a robust fea-

ture of normal somatic embryo development [

44 ]. Experiments

aimed at the separation of the sucrose osmotic inﬂ uence from its

role as carbon and energy source suggested that sucrose might

have an additional regulatory role in the maturation process. In P.

abies , a medium with 7.5 % PEG and 3 % maltose promoted the

development of a large number of somatic embryos, but with low

germination frequency in spite of the post-maturation partial des-

iccation [

45 ]. Conversely, somatic embryos developed on a

medium with 3 % sucrose (without additional osmotic agent),

3.3.3 Carbohydrates ,

PEG (Medium Water

Potential), and Gel Strength

(Water Availability)

Conifer Somatic Embryogenesis

140

although low in numbers, were able to germinate. A combination

of sugar assays, metabolic and proteomic analyses revealed that

somatic embryos grown on sucrose medium contained high levels

of sucrose, rafﬁ nose, and late embryogenesis abundant proteins, all

involved in the acquisition of desiccation tolerance (reviewed by

Trontin et al., Chapter

8 ). These embryos also accumulated starch

whereas those from PEG and maltose medium had high levels of

storage proteins. Therefore the poor germination of P. abies

somatic embryos grown on PEG and maltose medium was most

likely caused by the reduced desiccation tolerance.

Manipulation of water availability to the cells of EM was also

achieved by physical means, without affecting water potential of

the medium, by increasing the amount of gelling agent that

increased medium gel strength and consequently reduced the

amount of water available to the cells [

16 , 46 ]. By applying this

method of water control to P. strobus early embryo cultures, high

quality mature somatic embryo s were produced on medium with

1 % gellan gum that were characterized by a lower water content

compared with those from 0.4 % gellan gum medium. Combination

of 0.8 or 1 % gellan gum with 6 % sucrose (instead of 3 %) in the

maturation medium was even more beneﬁ cial because the somatic

embryos accumulated higher quantities of storage proteins [

47 ].

An intuitive interpretation of these results is that the developing

embryos must have been exposed to the water stress (drought type

of conditions) on the media with high gelling agent concentra-

tions, a condition similar to that of a developing zygotic embryo ,

when the maturation of the embryo is accompanied by desiccation

(loss of water). However, in a later study involving cultures of

Larix x eurolepis , an opposite conclusion was reached [

48 ]. The

more numerous and higher quality (lower water content) somatic

embryos that developed on medium with 0.8 % gellan gum were in

fact less stressed than those developed on 0.4 % gellan gum. This

conclusion was based on the measurements of physiological param-

eters and on the two-dimensional (2-D) protein gels that identiﬁ ed

62 proteins that differed between the two somatic embryo groups

from the two treatments. Fifty six proteins were subsequently iden-

tiﬁ ed, and among them 6-phosphogluconate dehydrogenase

(decarboxylating), actin, enolase, fructose phosphate aldolase,

phosphoglucomutase, and superoxide dismutase, which are known

to be associated with water stress, were expressed at a higher level

in somatic embryos developing on medium with 0.4 % gellan gum.

In addition to the increased abundance of heat shock proteins in

somatic embryos cultured on the 0.4 % gellan gum medium, the

observed increases in expression of pyruvate decarboxylase (which

directs carbon metabolism toward glycolysis) and apparent detoxi-

ﬁ cation capacity (indicated by the increased expression of superox-

ide dismutase) suggested that maturation medium containing 0.4 %

gellan gum induced a water stress response in the developing

somatic embryos. Contrary to this, somatic embryos developed on

Krystyna Klimaszewska et al.

141

0.8 % gellan gum medium accumulated stress proteins at a much

lower level. Further evidence supporting utilization of high gellan

gum medium for the maturation of somatic embryos came from a

large study of P. pinaster where multi-scale integrated analysis was

used to follow early molecular and physiological events involved in

somatic embryo development [

49 ]. Similarly to P. strobus , early

somatic embryos of P. pinaster do not develop on medium with

0.4 % gellan gum; instead, abundant proliferation of EM occurs,

which is not conducive to subsequent embryo development.

According to the transcriptomic and proteomic analysis results,

these cultures had enhanced glycolysis whereas those from medium

with 0.9 % gellan gum had adaptive, ABA -mediated molecular and

physiological responses marked by active protein synthesis and

overexpression of proteins involved in cell division, embryogenesis

and starch synthesis. Concomitantly, synthesis of protective sec-

ondary metabolites and regulation of oxidative stress were acti-

vated, most likely to adapt to the culture conditions. Furthermore,

two genotypes of cotyledonary P. pinaster somatic embryos, after

10–14 weeks of culture on maturation medium, were compared

with zygotic embryos excised from developing fresh and desic-

cated seeds with respect to dry mass, water content, sucrose and

rafﬁ nose contents, rafﬁ nose/sucrose ratio, and total proteins ([

50 ];

reviewed by Trontin et al., Chapter

8 ). The study demonstrated

that somatic embryos were the most similar to zygotic embryos in

seeds collected from late July to early August, and with respect to

total protein content up to October (Northern Hemisphere). The

somatic embryos, which typically are harvested after 12 weeks, are

not at the same maturity level as their zygotic counterparts at the

mature, desiccated stage.

There is evidence suggesting that ethylene , a gaseous plant growth

hormone produced by cultured plant cells and tissues, may also

affect the development of somatic embryo s in P. glauca [

51 ]. Due

to the volatile nature of ethylene, it is difﬁ cult to control it in cul-

ture. However, the reduction in endogenous ethylene synthesis was

achieved by incorporating α-aminooxyamino acid (AOA), a potent

inhibitor of ethylene biosynthesis, into the medium, which proved

to be beneﬁ cial to somatic embryo development . The mechanism

of this stimulation was not elucidated, only suggesting that AOA

may have interfered with the metabolism of other compounds,

most likely through the availability of S-adenosylmethionine, a

common precursor for both ethylene and polyamine biosynthesis.

In P. mariana , it has been shown that limiting ethylene biosyn-

thesis or its physiological action was beneﬁ cial to somatic embryo

development in a poor line, but not beneﬁ cial in a line that was a

good embryo producer [

52 ]. These opposite reactions could stem

from different initial ethylene levels in the two cultures, one hav-

ing a super-optimal and the other an optimal level. Later study

with cultures of P. sylvestris conﬁ rmed that ethylene production

3.3.4 Ethylene

Conifer Somatic Embryogenesis

142

varied among ﬁ ve embryogenic lines, both when cultured on

proliferation medium with 2,4-D and on maturation medium,

indicating a lack of deﬁ nite trend [

53 ]. Future experiments should

include many more genotypes to realize the full impact of ethyl-

ene on SE in P. sylvestris . Also, any generalization to other conifer

species should be avoided.

Among Abies species, A. nordmanniana somatic embryo s were

stimulated to develop on maturation medium (with ABA ) after a

4- to 8-week treatment with PCIB, 2-( p -chlorophenoxy)2-

methylpropionic acid, an auxin antagonist that is believed to reduce

the activity of endogenous indole-3-acetic acid (IAA) by competi-

tive binding to auxin receptors [

54 ]. Abies species do not require an

auxin for initiation or proliferation of early somatic embryos and

the subsequent problems pertaining to somatic embryo develop-

ment have been attributed to the high activity of endogenous auxin,

at least in A. nordmanniana . Treatment of P. sylvestris early somatic

embryos with the auxin transport inhibitor 1- N -naphtylphthalamic

acid (NPA) caused the embryos to form supernumerary suspensor

cells at high frequency, which led to abnormal development [

36 ].

Although treatment with PCIB increased the yield of somatic

embryos, their morphology was not affected, suggesting that the

supernumerary suspensor cells in early somatic embryos were stim-

ulated by disturbed polar auxin transport.

P. glauca somatic embryo development and maturation were

greatly improved through the manipulation of glutathione redox

status in EM cultures. By employing a two-step protocol that first

included reduced glutathione (GSH) and then its oxidized form

(GSSG) in culture medium, which caused a shift in the total glu-

tathione pool towards its oxidized state, proper somatic embryo

development was achieved [

42 ]. However, due to the high cost

and labor associated with this protocol, a simpler alternative was

developed involving dl - buthionine -[ S , R ]- sulfoximine (BSO)

[

55 ]. BSO is effective in reducing endogenous GSH levels through

the inhibition of its de novo synthesis without affecting glu-

tathione reductase, the GSH-recycling enzyme. These changes are

similar to those observed when GSH and GSSG are applied

sequentially to impose an oxidized environment. To maximize

somatic embryo development, BSO concentration had to be at

0.01 mM, while higher concentrations were inhibitory. Therefore,

it appears that certain threshold of cellular GSH must be main-

tained for embryo development to continue. In Araucaria angus-

tifolia cultures of EM, manipulation of the GSH/GSSG ratio of

the culture medium proved to be beneficial to somatic embryo

development up to the pre-cotyledonary stage, but to achieve a

complete development would require further modification of the

redox potential of the cultures [

56 ].

3.3.5 Antiauxin

and Auxin Transport

Inhibitor

3.3.6 Redox Compounds

Krystyna Klimaszewska et al.

143

When Larix laricina early somatic embryo s were cultured in high

density suspension in a liquid medium the differentiation of sus-

pensors was inhibited, thus negatively impacting the development

of new somatic embryos [

57 ]. It was conﬁ rmed that an inhibitory

compound was present in the conditioned culture medium, which

was subsequently puriﬁ ed and the compound was identiﬁ ed as

vanillyl benzyl ether (VBE) [

34 ]. Tests with synthetic VBE in the

medium produced similar results. Interestingly, the low density

suspension cultures also contained VBE, but at much lower con-

centrations, which did not prevent differentiation of the suspen-

sors. This ﬁ nding emphasizes the importance of the presence of

suspensors in somatic embryo development of a conifer and

increases awareness of the inﬂ uence of cell density on the embryo-

genic characteristics of a culture. Another modiﬁ er of normal

somatic embryo development is an inhibitor of polar auxin trans-

port, 1- N -naphtylphthalamic acid (NPA), which was tested in P.

abies [

35 ]. Polar auxin transport is essential to proper embryo pat-

terning and establishment of root/shoot polarity . During early

somatic embryo development, treatment with NPA caused an

increase in IAA content, abnormal cell division, and decreased pro-

grammed cell death resulting in the aberrant development of

embryonal tube cells and suspensors. These embryos had abnor-

mal morphology marked by malformed and fused cotyledons and

irregular cell divisions at the site of root meristem.

4 Extending Somatic Embryogenesis Protocols to Numerous Conifer Species

Previously published protocols have been utilized, often with slight

modiﬁ cations and with various degrees of success, to test/achieve

SE in numerous other conifer species.

Among Picea species, results that showed regenerated somatic

seedlings were published for P. morrisonicola [

58 ], P. koraiensis

[

59 ], and P. likiangensis [ 60 ]; however, it is not clear whether any

of the somatic trees were established in the ﬁ eld.

In the Pinus genus, at least 16 new species were reported to

display SE; however, for many of them, initiation and/or maturation

efﬁ ciencies and plant regeneration were low and needed further

research and improvements. Notoriously low initiation and survival

of EM has been reported for P. contorta [

61 ], P. monticola [ 29 ],

P. roxburghii [

62 , 63 ], P. pinaea [ 64 ], P. banksiana [ 65 ], P. densi-

ﬂ ora [

66 , 67 ], P. rigida x taeda [ 68 ], P. kesiya [ 69 ], P. thunbergii

[

70 ], P. armandii [ 71 ], and P. luchuensis [ 72 ]. On the other hand,

species such as P. patula [

73 ], P. nigra [ 22 ], P. bungeana [ 74 ],

P. brutia [

75 ], P. oocarpa [ 76 ], and P. halepensis [ 77 ] responded at

frequencies ranging from 9 to 30 %.

3.3.7 Inhibitors of SE

4.1 Pinaceae

Conifer Somatic Embryogenesis

144

Signiﬁ cant progress in SE response and plant regeneration has

been achieved for Larix leptolepis [

78 , 79 ], L . x eurolepis and L. x

marschlinsii [

80 ]. The reciprocal hybrids between L. decidua and

L. leptolepis are important species in Europe, and the ﬁ rst hybrid

variety (‘REVE-VERT’) was registered in France in 2005. SE

modiﬁ ed protocols were subsequently tested as a means for rapid

cloning of limited numbers of hybrid seeds and resulted in up to

48 % of initial zygotic embryo s producing high numbers of vigor-

ous somatic plants (Fig.

1 ). It is anticipated that SE will inﬂ uence

breeding strategies for these hybrids by offering an additional tool

for the production of large quantities of plants for clonal ﬁ eld tests.

SE in Abies species has been very challenging, but recent prog-

ress made with some species is encouraging. A study with A. alba

by Krajňáková et al. [

81 ] tested several variables for maturation of

somatic embryo s, which improved the maturation yield by utiliz-

ing a method of spreading the cultures in a layer on Whatman #2

ﬁ lter paper placed on the surface of a semisolid medium [

82 ] sup-

plemented with 32 μM ABA , maltose , organic N additives, and

devoid of PEG . The medium formulation was Murashige and

Skoog [

83 ] modiﬁ ed by 50 % reduction in inorganic salt strength.

The somatic embryos required desiccation for proper germination

Fig. 1 Larix x eurolepis “Reve-Vert” somatic seedlings (INRA, France, improved variety) acclimatized at

the XYLOBIOTECH nursery (XYLOFOREST platform, www.xyloforest.org ) located at the FCBA, Pierroton,

France (0.8×)

Krystyna Klimaszewska et al.

145

but still the conversion to plants remained inefﬁ cient. In A. lasiocarpa ,

a novel medium was designed based on the elemental analysis of

megagametophytes and designated AL, which was free of inor-

ganic nitrogen and in which

L -glutamine was supplied at 2 g/L as

the sole nitrogen source [

84 ]. AL medium was compared with

Schenk and Hildebrand [

85 ], and although initiation of SE (up to

37 %) did not differ signiﬁ cantly between the two media, EM

growth after subculture and its survival was better on AL. However,

the conversion to plants was very low (approximately 8 % of germi-

nated somatic embryos), which seems to be a norm in this genus.

Similarly, in A. cilicica and A. cilicica x A. nordmaniana , relatively

high initiations of EM were obtained (63 and 28 %, respectively)

but only a third of EM lines developed cotyledonary stage somatic

embryos on both maltose or lactose media [

86 ]. In A. numidica ,

both the maturation and germination of somatic embryos were

studied. PEG and 6 % maltose in maturation medium were very

effective followed by partial desiccation and germination of somatic

embryos on medium with activated charcoal and indolbutyric acid

(IBA) [

87 , 88 ]. In A. cephalonica , up to 25 % of the seeds pro-

duced EM lines, and somatic embryo development was achieved

on medium with PEG and sucrose followed by medium without

PEG for up to 12 weeks [

89 , 90 ]. However, germination was poor,

most likely due to the omission of partial desiccation of somatic

embryos before germination, which appears to be a requisite in

this genus.

A complete protocol for SE and plant production, including

cryopreservation, was reported recently for Tsuga caroliniana and

T. canadensis [

91 ]. Induction frequencies were from 17 to 52 % for

immature embryos , respectively. The results conﬁ rmed the inter-

play among the collection date of the cones, medium composition

and source tree on the frequency of SE induction, which has been

reported in all previous publications. Maturation of somatic

embryo s was completed by slow drying under permeable plastic

ﬁ lm. However, conversion of somatic embryos to plants was very

low and requires further research.

Cryptomeria japonica SE was successful with up to 17 % of imma-

ture seed explants tested over 3 years, and the presence of PGRs

was not required in the initiation medium [

92 ]. The embryogenic

characteristics of the cultures could be improved by increased glu-

tamine concentration in the medium [

30 ]. Somatic embryo s devel-

oped better on a medium that in addition to ABA , PEG , charcoal

and maltose , also contained 32 nM phytosulfokine [

32 ].

Two species of Chamaecyparis produced plants through SE,

namely C. pisifera [

93 ] and C. obtusa [ 94 , 95 ]. Initiation of SE

occurred on both a medium with PGRs (2,4-D and BA) and on

PGR -free medium at the frequency up to 33 and 48 %, respec-

tively. Development of somatic embryo s was promoted by PEG ,

4.2 Cupressaceae

Conifer Somatic Embryogenesis

146

AC , ABA , and maltose in a medium gelled with 0.3 or 0.5 % gellan

gum. Sixty-eight percent of C. obtusa EM lines produced cotyle-

donary somatic embryos, 91 % of which germinated and the plants

were subsequently transferred to a greenhouse [

95 ]. In both spe-

cies, somatic embryo germination was very high and conversion to

plants was not problematic. Field tests of C. pisifera somatic trees

are underway [

93 ].

Another species that was recently studied for its propensity to

undergo SE in the same family was Juniperus communis [

96 ]. This

work conﬁ rmed that similarly to Cryptomeria and Chamaecyparis ,

the presence of an auxin and/or a cytokinin was not necessary for

SE to be initiated and was even inhibitory in J. communis , with a

reduction from 50 to 25 %. Proliferation of EM was rapid on PGR-

free medium as well, but the maturation of somatic embryo s was

stimulated by 60 μM ABA after brief culture on ABA-free medium

with a lower concentration of N and Ca. The development of

somatic embryos was highly asynchronous and the culture pro-

duced a continuous supply of early and mature somatic embryos.

The latter germinated after partial desiccation and converted to

plants at a low frequency. A major impediment to plant production

was the growth of new EM at the basal part of the somatic plant,

which could not be controlled by the application of the gibberellin

(GA

4/7 ) in the medium.

In Taxus wallichiana , SE was achieved indirectly from calli that

grew on zygotic embryo s in culture [

97 ]. Initially, the explants

were cultured in the presence of BA and NAA, and after 8 weeks

they produced compact yellow calli. Subsequently, when trans-

ferred onto a medium with 2,4-D, NAA, and BA, the calli changed

in morphology, and two out of four displayed embryogenic char-

acteristics. Somatic embryo development was achieved on medium

with ABA and charcoal after 12 weeks; however, the conversion

rate to plants was only 10 %.

A complete SE protocol was developed for Torreya taxifolia [

98 ].

High initiation of SE (60–100 %) from six seed families was accom-

plished on medium with various additives, including PGRs (2,4-D,

BA, kinetin, and ABA ) and maltose . The EMs were cryopreserved

using the standard protocol. Somatic embryo s developed on a

medium with ABA, activated charcoal , maltose, biotin, brassino-

lide, MES, folic acid, and pyruvic acid. Two genotypes of clonal

mature somatic embryo s germinated at 64 and 95 %, respectively,

and after 2 months the somatic seedlings were planted in a sub-

strate. The species is under threat of extinction and SE will assist in

the present and future conservation of this ancient plant.

Despite extensive research to develop SE protocols for Araucaria

angustifolia , only pre-cotyledonary somatic embryo s were obtained

in culture [

99 , 100 ].

4.3 Taxaceae

4.4 Cephalotaxaceae

4.5 Araucariaceae

Krystyna Klimaszewska et al.

147

5 Genetic Stability of Cultured Embryonal Mass and Somatic Seedlings

Embryogenic cell lines from ten half-sib seed families of P. sylvestris

were analyzed using four nuclear single sequence repeat (SSR)

markers, also known as microsatellites or short tandem repeats

(STR) [

101 ]. The aim was to determine whether the genetic stabil-

ity of the lines changes during in vitro culture. The results indicated

that mutations occurred in the cell lines and that their frequency

was dependent on the seed family. Interestingly, mutations were

detected in cell lines cultured on both a medium with PGRs and on

a medium without PGRs, undermining the present notion that

2,4-D is the main culprit causing genetic instability. The authors

also found a considerable mutation rate in zygotic embryo s, albeit

at a much lower rate compared with cultured EM , which led them

to conclude that the in vitro culture stress triggered the mutations

in the microsatellite regions in P. sylvestris . Whether the instability in

the studied microsatellite loci reﬂ ect alteration in functional genes

remains to be investigated. In a similar study with P. pinaster , 17

EM lines from six seed families were analyzed using seven nuclear

SSR markers after 6, 14 and 22 months of culture as well as regen-

erated somatic seedlings [

102 ]. The SSR pattern at the time of line

establishment was used as a reference for the cultures of increasing

age. Genetic variation was detected in cultures of all ages and in 5

out of 52 somatic seedlings. Some somatic seedlings displayed pla-

giotropism and loss of apical shoot dominance, but no correlation

was found between genetic instability at the analyzed loci and the

abnormal phenotype. Nevertheless, there is a risk of genetic muta-

tions during the cell proliferation stage in vitro, which may lead to

the regeneration of mutant plants with different mutations occur-

ring among somatic plants regenerated even from the same EM

line. The latter could be caused by the presence of a mixture of cells

that accumulated different mutations in a given cell line and, hence,

the regenerated plants were not clonal. There is some speculation

that the loss of embryo development capacity in aged EM lines

might be attributed to the accumulation of mutations, perhaps

together with epigenetic changes, during prolonged in vitro cul-

ture, as described below.

6 Aging of Embryogenic Cultures and its Inﬂ uence on the Ability to Produce

Mature Somatic Embryos

The age of embryogenic cultures maintained on semisolid medium

can negatively inﬂ uence their ability to produce mature somatic

embryo s in some conifer species. However, aging was not counter-

productive in cultures of Picea and Larix hybrids. In L. x eurolepis ,

a line was still productive after 9 years of subculturing [

80 ]. On the

contrary, the somatic embryo regeneration ability of cultures of

Conifer Somatic Embryogenesis

148

A. lasiocarpa [ 84 ] and C. japonica [ 92 ] decreased overtime

whereas in Larix leptolepis , embryogenic cultures became non-

embryogenic [

103 ]. The aging and associated changes appear

critical in Pinus sp. In P. pinaster , maturation yield decreased rap-

idly within 6 months of culture [

28 , 104 ]. Reduction in the quantity

of somatic embryos or cessation of somatic embryo development

was accompanied by substantial modiﬁ cations to the cellular orga-

nization/composition of the culture during proliferation [

27 ].

Total culture time also affected the quality of cotyledonary somatic

embryos, with progressive reduction of size and germination rate

[

28 ]. However, the embryogenic culture’s ability to regenerate

cotyledonary somatic embryos could be prolonged by modifying

the culture medium composition and subculture frequency [

27 ].

In Larix sp., the effect of aging on embryogenic ability has

been studied at the molecular level (reviewed by Trontin et al.,

Chapter

8 ). Differential expression of various microRNAs (four

major miRNA families: miR171, miR159, miR169, miR172) has

been detected in embryogenic and in non-embryogenic cultures of

Larix kaempferi [

105 ]. In particular, miR171 and miR159 were

found downregulated and upregulated in non-embryogenic cul-

tures, respectively. Subsequently, the authors identiﬁ ed a MYB

transcription factor ( LaMYB33 from L. kaempferi ) as a target gene

for miR159 and Larix SCARECROW-LIKE 6 homolog ( LaSCL6 )

was targeted by miR171 [

106 ]. Post-transcriptional regulation of

LaMYB33 and LaSCL6 by miRNAs may participate in the mainte-

nance of embryogenic potential as part of the epigenetic complex

of regulation of gene expression [

103 , 106 ].

To circumvent the recurrent problem of aging, the embryo-

genic cultures of most conifer species are routinely cryopreserved

shortly after initiation ( see Ozudogru and Lambardi, Chapter

32 ).

Full embryogenic competence of old embryogenic lines could be

restored in P. pinaster by inducing new cultures from cotyledonary

somatic embryo s (secondary SE; [

104 ]).

7 Desiccation and Cryopreservation

The ability to cryopreserve EM has been a critical success feature

of the development of SE to the level we utilize this technology in

the afforestation programs today. The maturation competences of

SE cultures under conditions of continuous subculture are highly

variable both within and between species. Some species of Picea

and Larix are seemingly unaffected by long periods of minimal or

erratic subculture, but many Pinus species show a sharp decline in

both quantitative and qualitative mature embryo production [

28 ,

80 , 107 ]. These differences, to some degree, inﬂ uence the urgency

to cryopreserve cultures. However, the cost savings associated with

not subculturing and the importance of retention of genetic ﬁ delity

Krystyna Klimaszewska et al.

149

and maturation competence make cryopreservation as critical as

the other stages of the SE process.

Considering the progress made in cryopreservation over the

past decade, key developments include a better understanding of

the morphology of cell lines, the interaction of cryoprotectant

treatments with these and the development of protocols that facili-

tate storage of immature and mature somatic embryo s [

108 , 109 ].

It is also clear that relatively simple pretreatments and freezing pro-

tocols are proving as effective as earlier more complex methodolo-

gies. Embryogenic cell lines on proliferation media are highly

heterogeneous, consisting of a range of cell types in a state of con-

stant differentiation and dedifferentiation. This variability in cell

types has been thought to differentially inﬂ uence the responsive-

ness of cellular components to osmotic treatment, colligate cryo-

protection and controlled cooling [

107 ]. Recent results with a

number of species indicate that many of the factors that contribute

to successful cryopreservation still remain elusive [

108 ]. The

increasing body of work using more differentiated tissues may lead

to this being the preferential material for long-term storage of

conifers.

Cryotolerance of P. abies was studied in association with growth

rate, anatomical features and polyamines (putrescine, spermidine

and spermine) in ﬁ ve embryogenic cultures [

108 ]. The authors

found that the ability to produce normal mature embryos was the

only characteristic shown to have a positive correlation with cryo-

tolerance. Of the two lines showing a high percentage of cryotoler-

ance, one was a highly productive line, in terms of maturation

ability, the other one had a negligible ability to produce mature

embryos. Anatomically, the contrast between the embryo initials

for these two lines prior to cryopreservation was striking, with the

poor-embryo-producing line showing highly dedifferentiated ini-

tials. The same contrast was seen with total polyamine contents,

with the two cell lines with the highest contents giving opposite

results with regard to cryotolerance at 94 and 0 %. These observa-

tions indicate that the factors that confer cryotolerance in EM are

yet to be fully elucidated.

New species to show successful recovery from liquid nitrogen

storage include P. nigra and P. omorika [

110 – 112 ], respectively.

More unusually for conifer embryogenic tissues, the pretreatment

stages for P. omorika were done on semisolid medium with increas-

ing sucrose concentrations followed by air drying of the EM to

20 % of original fresh weight and subsequent immersion directly

into liquid nitrogen. No other cryoprotectant agents were used.

After cryostorage, P. nigra demonstrated growth rates and ability to

produce mature embryos similar to the control material maintained

in long-term culture [

110 ]. Another less common pretreatment

( maltose ) and cryoprotectant formulation was applied to P. pinaster

7.1 Cryotolerance of

Embryonal Masses

Conifer Somatic Embryogenesis

150

that included PEG 4000 with dimethylsulfoxide ( DMSO ), resulting

in 97 % recovery of the cell lines tested [

113 ].

Vitriﬁ cation using a modiﬁ ed plant vitriﬁ cation solution

( PVS2 ), developed primarily for nonembryogenic tissues and shoot

apices, has been tested with the aim of developing a simpliﬁ ed cryo-

preservation procedure for conifer embryogenic tissues. Successful

vitriﬁ cation of tissues would facilitate immediate immersion into

liquid nitrogen storage without intervening steps including tran-

sient storage at −40 to −80 °C in freezers or programmable cooling

incubators. This was successfully achieved with some cell lines of P.

mariana [

114 ]. An encapsulation /dehydration method was tested

with immature somatic embryo s of P. sitchensis and resulted in the

regeneration of EM following immersion in liquid nitrogen [

115 ].

No −40 to −80 °C or programmable freezer are required, but the

tissue treatment is labor intensive prior to storage.

A novel method for tissue regrowth was tested with P. radiata

and resulted in signiﬁ cantly improved post-thaw growth with 60

cell lines stored from 6 months to 4 years prior to thawing [

116 ].

The authors used a vigorous culture (nurse culture) of P. radiata

to nurse the thawed cells; the nurse culture and thawed cells were

separated from each other by a nylon screen. Further simpliﬁ cation

of methods was achieved with P. glauca x engelmannii and P. men-

ziesii . The method eliminated both the use of toxic cryoprotec-

tants and freezing environments. Following culture on ABA

medium at 4 °C, the tissue was immersed directly into liquid nitro-

gen [

109 ]. The method relied on preconditioning of early somatic

embryo s and these retained the ability to regenerate EM following

storage in liquid nitrogen.

Contamination of EM lines can still plague this step of the SE

process, with a number of authors reporting signiﬁ cant losses of

cell lines upon thawing from liquid nitrogen storage [

108 , 112 ].

Picea omorika cryopreserved as clumps of tissues rather than as

cell suspension s had a decreased frequency of contamination if

liquid nitrogen was prevented from entering the vials during

freezing [

112 ]. In embryogenic cultures of P. radiata cell lines

stored for 6 months, none of the 37 genotypes (222 vials) were

contaminated and only 5 % of the vials from a further 23 geno-

types (138 vials) stored for 4 years were contaminated, despite the

fact that all vials had been immersed in liquid nitrogen upon freez-

ing [

116 ]. Interestingly, antibiotic cephotaxime (100 mg/L) was

used in the proliferation medium to reduce the risk of bacterial

contamination of cultures during the frequent treatments before

cryostorage of P. abies [

108 ].

Mature somatic embryo storage could potentially confer a range of

advantages over the cryopreservation of embryogenic masses and

would be especially useful in the application of this propagation

technology. Effective storage would facilitate both the synchrony

7.2 Mature Somatic

Embryo Storage

Krystyna Klimaszewska et al.

151

of seed orchard and laboratory production with seasonal nursery

and planting programs. Added advantages are that with careful

pretreatment, no cryoprotectant chemicals or programmable freez-

ing equipment is required. Successful desiccation without cryo-

preservation may also be an important aspect of improving quality

and synchronizing germination in somatic embryos in many

species. Continued development of direct sowing and artiﬁ cial

seed technologies may also beneﬁ t from more effective desiccation

protocols.

The desiccation environment seems to be one of the key ele-

ments for successful storage. Picea mariana and P. glauca somatic

embryo s were slowly dried at 97 or 88 % relative humidity (RH) to

reach a water content of 0.23 H

2 O g/L dry weight before

achieving a high post liquid nitrogen germination frequency of

93.8 % [

117 ]. Desiccation at a lower RH of 63 % had a signiﬁ cantly

negative effect on subsequent germination following cryopreserva-

tion. Interestingly, the somatic embryos that managed to survive

this treatment showed a 100 % conversion to plantlets whereas the

conversion of the somatic embryos from the 97 or 88 % RH treat-

ments ranged from 26.7 to 46.7 %. These authors also tested the

stored embryo potential for embryogenic tissue reinduction fol-

lowing thawing. Embryos that had been desiccated at high RH

(97 %) and were rehydrated for 12 h at 100 % RH had reinduction

rates that were similar to those of the controls [

117 ]. Further work

from this team has elucidated some of the mechanisms linked to

fast desiccation tolerance in P. mariana [

118 ]. Their studies

showed that an initial short period of slow desiccation of the

embryos increased their subsequent tolerance to a fast desiccation

treatment. The mechanisms behind this indicated an increase in

sucrose accumulation, occurrence of rafﬁ nose, and depletion of

starch reserves within the somatic embryos. The occurrence of

dehydrins was also investigated in reference to their suspected role

in the development of desiccation tolerance. The authors noted a

doubling of the dehydrin signal intensity after 48 h of slow desic-

cation (24–48 kDA), which coincided with the best treatment for

subsequent germination of rapidly desiccated embryos.

Picea glauca and P. glauca × engelmannii complex somatic

embryo s were gradually dried over salt solutions to the level of dry

seed embryos and retained their viability upon rehydration [

119 ].

Desiccated somatic embryos also survived subsequent freezing in

liquid nitrogen , without the addition of cryoprotectant or pre-

culture steps. Highest survival (>80 %) after freezing in liquid

nitrogen was in embryos pre-dried to Ψ of –15 to –20 MPa, which

yielded relative water content (RWC) close to predicted bound

(apoplastic) water values. In another study, somatic embryos of

P. glauca survived a rapid desiccation treatment (2 h of air drying

on a laminar ﬂ ow bench at ambient temperature and humidity) if

they were carefully preconditioned on maturation medium [

120 ].

Conifer Somatic Embryogenesis

152

The optimum treatment was leaving embryogenic tissue on

maturation medium for 51 days, making it possible for embryos to

become cotyledonary before placing the Petri dishes into 5 °C for

8 more weeks of incubation. It should be noted that in P. glauca ,

cotyledonary embryos are fully developed after 51 days and preco-

cious germination was observed in some embryos prior to incuba-

tion at 5 °C. In contrast to the results presented for P. mariana

(and P. glauca ) [

118 ], shorter periods of incubation were detri-

mental to the quality of the germinant following rapid desiccation

[

120 ]. Elucidation of the mechanisms behind the cold tolerance of

P. glauca somatic embryos has subsequently shown, with freezing

damage tests based on electrolyte leakage, that somatic embryos

matured at lower temperatures possessed signiﬁ cantly higher freez-

ing tolerances than somatic embryos matured at 20 °C [

121 ].

8 SE from Vegetative Tissues of Adult Conifers

Vegetative (also known as clonal) propagation of adult trees has a

major advantage over propagation through seed because large

genetic gains are achieved by capturing a large proportion of tree

genetic diversity in a single selection cycle [

122 ]. Hence, vegeta-

tive propagation of select superior forest conifers through SE is

highly desirable, particularly because it has the potential to deliver

a stable supply of superior seedlings for forest plantations. However,

in spite of decades of research efforts, efﬁ cient propagation of adult

conifers by any means is still beyond reach [

61 , 123 , 124 ]. The

ﬁ rst work that raised expectations in this area of research was induc-

tion of SE in buds (primordial shoots/needles) of 2- to 3-year-old

P. abies grown from a somatic embryo [

125 ] and Ceratozamia spp.

[

126 ], but no results on somatic plant growth have been pub-

lished. Using four genotypes of somatic trees of P. glauca , it was

subsequently demonstrated that one genotype produced SE from

primordial shoot explants (Fig.

2a–f ) consistently from age 2 (in

2002; [

127 ]) to 15 years (in 2015; Klimaszewska, personal com-

munication). The media for each stage of SE were the same as

those used for seed embryo SE and a large number of juvenile

propagules (somatic seedlings) have been grown in a greenhouse

and subsequently planted in the ﬁ eld. These somatic trees derived

from donor trees of increasing chronological and ontogenic ages

Fig. 2 (continued) (17.5×). ( c ) Primordial shoot cut longitudinally and showing slightly elongated needle

primordia (24×).Fig. 2 (continued) ( d ) Embryonal masses growing from the explant after 33 days (magniﬁ cation

25×). ( e ) Mature somatic embryos produced from embryonal masses induced from the same donor trees of

different ages; 2, 7, and 8 years old (0.5×). ( f ) Somatic seedlings cultured on germination medium for 7 weeks

(1.1×). ( g ) Clonal, juvenile G6 trees produced from primordial shoots collected from 7-year-old (on the left ) and

8-year-old (on the right ) donor trees (1×) growing in the nursery of NRCan-CFS, Valcartier, QC, Canada

Krystyna Klimaszewska et al.

Fig. 2 Induction of somatic embryo genesis (SE) within primordial shoots of somatic Picea glauca 13-year-old

trees, genotype G6. Vegetative buds were collected on May 6, 2013 from a plantation established in 2003 by

NRCan-CFS in Valcartier, QC, Canada. ( a ) A branch with pre-ﬂ ush buds (2.5×). ( b ) Cleaned and disinfected buds

154

are being evaluated for their growth rate and morphology, and are

expected to provide evidence of true rejuvenation. Simultaneously,

the donor trees of responsive and nonresponsive genotypes pro-

vided a unique opportunity to examine the molecular aspects

underpinning SE within shoot tissues of adult P. glauca trees

(reviewed by Trontin et al., Chapter

8 ). A 32,000 oligo-probe

microarray was used for transcriptome-wide expression proﬁ ling of

explants at day 0 and day 7 of culture, which led to the identiﬁ ca-

tion of four of the most differentially expressed genes in each of the

two genotypes [

128 ]. The absolute quantitative PCR (qPCR) of

these genes was expanded to 21 days of SE induction and showed

that the expression of all eight genes was maintained throughout

the induction period. In contrast to the responsive genotype,

explants of the nonresponsive genotype expressed high levels of

stress-related genes, such as two extracellular serine protease inhib-

itors, a cell wall invertase, and a class III apoplastic peroxidase,

whereas the former showed temperate expression of these genes.

Instead, high expression of dehydrins and the QT-repeat and pro-

line rich proteins that are conifer-speciﬁ c were identiﬁ ed in the

responsive genotype and suggested an adaptive stress response .

These results further suggested that the possible causes of the lack

of SE induction in an explant may not be necessarily due to an

innate lack of SE promoting activity, but that biotic defense activa-

tion could potentially be a dominant antagonist. Therefore, future

work should focus on determining how and if suppressing biotic

defense activation could be used to promote SE induction in non-

responsive explants.

9 Field Growth of SE Trees

Clonal forestry offers signiﬁ cant advantages for forest productivity

due to the genetic gain (volume and quality improvements) that

can be realized through selection and mass propagation of elite

individuals (clones) [

129 ]. Somatic embryo genesis, with its capac-

ity for long-term germplasm cryopreservation and scale-up tech-

nologies, is the preferred avenue to accelerate the selection and

operational deployment of value-added genotypes, especially

through multivarietal forestry [

4 , 7 ]. Over the past decade, more

information has become available from ﬁ eld performance trials of

planting stock derived from SE. As described in an earlier review,

the majority of reports was for Picea spp. due to their responsive-

ness to SE relative to other genera [

4 ]. A number of early trials

with Picea spp. and P. menziesii have been established for several

decades and while the Pinus spp. have been more recalcitrant to

SE, some information is available [

7 ].

Further studies looking at the possible long-term effects on

ﬁ eld growth of somatic seedlings caused by in vitro conditions

Krystyna Klimaszewska et al.

155

was undertaken with P. abies [ 130 ]. The somatic plants were

assessed for survival and early growth after 4 months in the ﬁ eld.

The authors conﬁ rmed that prolonged exposure to ABA during

the maturation period of somatic embryo formation inhibited early

growth. Another treatment of continuous light routinely given to

P. abies seedlings to improve early growth in the greenhouse had a

negative effect on the growth of somatic plants. The authors con-

cluded that direct inwintering of somatic plants after transfer to ex

vitro conditions should be avoided. Early greenhouse work study-

ing clonal variation in morphology, growth, physiology, anatomy,

and ultrastructure of 6-month-old container-grown P. glauca

somatic plants found a number of differences when compared with

zygotic seedlings of the same families [

131 ]. Height ranges of

clones were greater (14.4–31.8 cm) than that of seedlings (15.8–

24.3 cm), and root collar diameters were generally greater in

clones. Variation within families was larger among somatic clones

than among zygotic seedlings for height, needle dry mass and

branch density. Light microscopy showed that tannins were more

abundant in somatic plants than seedlings; otherwise all needle

samples displayed a similar morphology. Of more concern was

the incidence of root deformation in somatic plants which had to

be transplanted from culture vessels to styroblock containers.

Only 52 % of somatic plants had a normal root form, a rate that is

comparable with that observed in zygotic seedlings that were not

transplanted. Another interesting observation was that plants from

speciﬁ c clones suffered from copper deﬁ ciency symptoms in all rep-

lications despite fertilizer application. What was clear and encour-

aging from this study was the early screening potential for selection

of superior clones based on both physiological and morphological

characteristics [

131 ]. Subsequent work presenting pooled data

that compared zygotic seedlings and somatic plants of P. menziesii

for gas exchange rate s, water relations and frost hardiness after 2

years in the ﬁ eld concluded that there were no signiﬁ cant differ-

ences between the two stock types [

132 ]. However, no data was

presented for individual clone performance. There were only three

clones in this trial derived from control crosses versus the seedling

controls, which were from bulked open pollinated seed collected

from the same orchard. When considering frost hardiness and bud

break, no signiﬁ cant differences were found between the two stock

types in the latter study. More recent work raised the possibility

that there may be some interaction between temperature at the

time of somatic embryo maturation and subsequent frost hardi-

ness and bud break especially in the ﬁ rst few years of plant estab-

lishment [

117 ]. Based on ﬁ eld performance studies, it appears

that clones produced from SE, at least those of P. menziesii and

P. glauca , can be highly acclimated to different climatic conditions

[

133 , 134 ].

Conifer Somatic Embryogenesis

156

Evaluation of genetic parameters and examination of genotype

x environment interactions to characterize the genetic stability of

somatic seedlings of P. glauca have been done 4 years after estab-

lishment of the ﬁ eld tests [

134 ]. In these tests, 52 clones (from 14

control-crossed families) were compared and they are the ﬁ rst of a

series of trials established under different ecological site conditions

comparing over 1000 somatic clones. Encouragingly, the percent-

age of somatic seedlings (52 clones) exhibiting normal adaptive

characteristics for survival (98–99 %) and bud frost damage and

stem form (90–99 %) characteristics were high, and therefore,

genetic parameters were not calculated for these characteristics.

Strong positive genotypic correlations were found between height,

diameter, annual shoot length and volume. The authors felt that

the stability of the clonal performance at the two sites reﬂ ected the

efﬁ ciency of clonal selection and was therefore a good reason to

promote the selection of generalist clones for future applications in

multiclonal forestry [

134 ]. Older somatic plantings (5.5 years) of

P. menziesii var. menziesii have been assessed for survival and per-

formance, clonal genetic parameters such as variances, heritability,

and correlations, and for stability of clonal performance ac ross ﬁ ve

sites in Washington and Oregon, in the Paciﬁ c Northwest, USA

[

133 ]. There were 70 clones in the test and the somatic seedlings

were grown in the same greenhouse for 1 year prior to planting.

All exhibited growth rates and morphology within the normal

range exhibited by zygotic seedlings in nurseries. The survival of

the somatic seedling clones at 5.5 years ranged from 92 to 99 %

and the general conclusion from this study was that the stability of

these clones was encouraging for future clonal forestry applications

in coastal Douglas ﬁ r.

A set of P. radiata trials was established in New Zealand and

Australia (three in each country) to investigate clonal stability

focusing on growth and form traits [

135 ] (Fig. 3 ). The planting

stock was derived from cuttings taken from hedges established

from somatic embryo s rather than using germinated somatic

embryos directly. One reason for this approach was to improve

plant quality within clones (height, root mass, and stem diameter,

all of which were positively affected). There were 245–280 clones

tested at the three New Zealand trials and 44–69 clones at the

three Australian sites. In general, clonal stability was good ac ross

the New Zealand sites, and although there was only a small num-

ber of clones that were common between Australia and New

Zealand, clones stable for growth could be identiﬁ ed across both

countries. The authors did note that age 5 may still be too young

to draw ﬁ rm conclusions with regard to genotype rankings. Forest

Genetics Ltd. planted their ﬁ rst trials of P. radiata derived from

somatic seedlings in 1999. They have been able to clearly identify

outstanding clones, which now form the basis of ﬁ eld-proven

material being sold to commercial clients (

www.forest-genetics.com ).

Krystyna Klimaszewska et al.

157

These plants command a premium price relative to seedlings of

control pollinated seed lots. Evaluation of somatic seedlings has

been also ongoing in France since 1999 with P. pinaster for which ca.

3200 clonal trees from more than 200 genotypes were established

in eight ﬁ eld tests [

7 ]. Data analysis at age 6, from 24 clones

planted in 2004, indicated that somatic seedlings are producing

normal trees but usually with a lower initial growth rate than those

from seedlings (Trontin, personal communication). However, it

has been shown that mean relative increase in height was similar or

even higher in speciﬁ c somatic lines after 6 years, suggesting that

normal growth can be recovered later.

Seed production from somatic clonal trees has been studied in

P. mariana [

136 ]. The authors found that the somatic trees pro-

duced both viable pollen and female cones that were able to be

crossed to produce equally viable seeds. The authors noted that

male strobili were produced about 6 years after planting and 2

years after the early onset of female ﬂ ower production, which is

earlier than what is generally observed in zygotic trees. The

authors also noted the incidence of albino germinants (up to 14 %

in one particular inbred cross). No direct non-somatic clone con-

trols were used in this research to determine if earlier male and

female cone production had an adverse effect on vegetative

growth. Results were generally compared with other data available

for P. mariana and there were no outstanding anomalies (pollen

germination , seed mass, and morphophysiological standards for

planting stock). The authors concluded that the stock produced

Fig. 3 Somatic Pinus radiata in a ﬁ eld test of the Forest Genetics Ltd., New Zealand

Conifer Somatic Embryogenesis

158

through SE and selected for exceptional performance could be

used for subsequent seed production and would enhance gains

from multivarietal forestry.

10 Bioreactor /Scale-Up Studies

For commercial application of SE, laboratory-scale protocols must

be scaled up and fulﬁ ll several criteria such as production of high

quantities of uniform somatic embryo s at a given time and at a

reasonable cost per unit. This can be achieved by utilizing

bioreactors, which are amenable to automation and allow contin-

uous monitoring and control of growth conditions (agitation,

pH, oxygen , and carbon dioxide ), large volumes, and mainte-

nance of a homogeneous culture. In a study with P. sitchensis , two

EM lines were grown in bioreactors of different conﬁ gurations

( air-lift , bubble, stirred tank, and hanging stirrer bar) and com-

pared with shake ﬂ ask cultures [

137 ]. The bioreactors were 5, 2,

or 1 L in volume. Both lines exhibited larger increases in biomass

when grown in bioreactors, but one line proliferated as single

early somatic embryos while the other one formed large aggre-

gates of somatic embryos. Samples taken from all cultures were

transferred onto maturation medium with 40 μM ABA , 1 μM

IBA, 3 % sucrose , and 0.1 % activated charcoal . There were two

medium variants in Petri dishes: one semisolid (0.6 % agar ) and

another semisolid covered with 7 mL of liquid medium for sub-

merged culture. One line produced cotyledonary somatic embryos

at the highest number when proliferated in bubble bioreactor on

both variants of maturation medium. For the second line, the sub-

merged way of culture was unsuitable. The results suggested that

the bioreactor conﬁ guration, design, and operating conditions

must be adequately chosen to suit the physiological, metabolic,

and morphological characteristics of a line. For P. menziesii , a

large-scale somatic embryo production system was developed by

Weyerhauser Co. (WA, USA) [

138 ]. It involved growing the EM

in 1 L ﬂ asks in liquid medium on a rotary shaker in darkness, fol-

lowed by culture in perfusion bioreactors on liquid medium

soaked pads containing PEG , ABA, GA

4/7 , and charcoal for devel-

opment and maturation. The development medium was pumped

from the reservoir into the bioreactor until it made contact with

the lower surface of the pads. The medium was absorbed in the

pads by capillary action and, after a few hours, it was pumped out

to the reservoir. This was repeated at regular intervals until mature

cotyledonary embryos developed. Twenty to ﬁ fty cotyledonary

somatic embryos were produced from the initial 1 mL of settled

EM, but different genotypes showed variations in both the num-

ber and quality of mature somatic embryos. The somatic embryos

were cold-treated before germination . The authors concluded

Krystyna Klimaszewska et al.

159

that the solution for low-cost mass production of conifers was the

combination of liquid culture with bioreactors, automation tech-

nology, and manufactured seed delivery system.

11 Conclusions and Future Research

The multidiscipline approach that is taking place to ﬁ nd solutions

to some of the problems still facing large-scale production of conifer

trees through SE should prove fruitful in the future. SE research

has generated knowledge and protocols that can be immediately

applied from one species to another and have their utility veriﬁ ed.

For example, in Pinus species and other conifer genera where ini-

tiation of SE is proving problematic, the viability of crosses prior

to sampling needs to be considered. Following sampling for

embryogenesis, a further collection of cones should be made

when the seed is mature to verify that the seed is viable. It has

been shown with a range of conifers that embryos often form early

in development only to abort prior to becoming cotyledonary,

especially in situations where self- fertilization or hybridization

may be occurring. In some cases, SE may be a way of rescuing

these embryos that could be advantageous, especially with the

establishment of novel ﬁ rst generation hybrids. However, for gen-

eral protocol development, it is better to have known high viabil-

ity crosses to work with. Another confounding factor with protocol

development is the presence of the megagametophyte , which is

likely to have a confounding effect with regard to development of

an optimal induction medium. The megagametophyte tissue dies

as soon as it is dissected away from the seed coat and may start to

produce toxic leachates. Examination of the literature does show

mixed responses and it is possible that the megagametophyte acts

as a buffer against suboptimal media interactions, and that it even

provides some nutritive beneﬁ ts in the short term. It is recom-

mended that both methods of dissection be tested as part of the

matrix including media modiﬁ cations when protocols are being

developed or improved for conifer species. Optimization of culture

medium should beneﬁ t from using software tools for experimental

design, computation, and graphic visualization of multifactor

interactions that together inﬂ uence the culture productivity from

its onset in vitro to the plants in a greenhouse/nursery as demon-

strated for herbaceous species by Halloran et al. [

139 ] and Adelberg

et al. [

140 ].

We expect to see increased integration of somatic embryo gen-

esis within current nursery practices. In the laboratory, methods to

reduce plant production costs will continue to develop; these

include liquid culture and sorting mechanisms for mature somatic

embryos. Somatic embryo s of a number of conifer species are

amenable to organogenesis, and this step will be used to provide

Conifer Somatic Embryogenesis

160

plants for stool bed and subsequent cutting regeneration. Where

physiological age constraints limit the life of stool beds, liquid

nitrogen storage will ensure a continuous supply of juvenile stock

plants. Soft tissue manipulation robotics for medical procedures is

improving as well as visual multidimensional graphics, and this may

create new opportunities for automation of the SE process.

Acknowledgements

We thank Dr. J. Bonga (Natural Resources Canada-Canadian

Forest Service) for comments on the early draft of the manuscript

and Mrs. I. Lamarre (NRCan-CFS) for English editing. J.F.T. and

M.A.L.W. were supported by the French “Région Aquitaine”

(EMBRYO2011: 09012579-045) project and by Conseil Régional

de la “Région Centre” (EMBRYOME project, contract 33639),

respectively.

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_8, © Springer Science+Business Media New York 2016

Chapter 8

Molecular Aspects of Conifer Zygotic and Somatic

Embryo Development: A Review of Genome-Wide

Approaches and Recent Insights

Jean-François Trontin , Krystyna Klimaszewska , Alexandre Morel ,

Catherine Hargreaves , and Marie-Anne Lelu-Walter

Abstract

Genome-wide proﬁ ling (transcriptomics, proteomics, metabolomics) is providing unprecedented opportu-

nities to unravel the complexity of coordinated gene expression during embryo development in trees, espe-

cially conifer species harboring “giga-genome.” This knowledge should be critical for the efﬁ cient delivery

of improved varieties through seeds and/or somatic embryos in ﬂ uctuating markets and to cope with climate

change. We reviewed “omics” as well as targeted gene expression studies during both somatic and zygotic

embryo development in conifers and tentatively puzzled over the critical processes and genes involved at the

speciﬁ c developmental and transition stages. Current limitations to the interpretation of these large datasets

are going to be lifted through the ongoing development of comprehensive genome resources in conifers.

Nevertheless omics already conﬁ rmed that master regulators (e.g., transcription and epigenetic factors) play

central roles. As in model angiosperms, the molecular regulation from early to late embryogenesis may

mainly arise from spatiotemporal modulation of auxin-, gibberellin-, and abscisic acid- mediated responses.

Omics also showed the potential for the development of tools to assess the progress of embryo development

or to build genotype-independent, predictive models of embryogenesis-speciﬁ c characteristics.

Key words Developmental regulator , Embryo patterning , Gymnosperm , Metabolome , Proteome ,

Somatic embryo genesis , Transcriptome , Stress

1 Introduction

Compared with herbaceous angiosperms such as Arabidopsis thali-

ana , elucidation of the molecular events regulating embryo devel-

opment in trees, and particularly in conifers (the primary source

for wood production worldwide), has been hindered by their large

physical size, slow growth, long generation time, and very large

genome. A number of powerful genetic approaches whose efﬁ -

ciency does not require the availability of large genomic resources

or full genome sequence (e.g., embryo defective mutants, T-DNA

insertional mutagenesis) are therefore impracticable with conifers

168

[ 1 , 2 ]. The recent implementation of qualitative and quantitative

methods for the genome-wide proﬁ ling of genes [

3 ] (transcrip-

tomics), proteins [

4 ] (proteomics), and metabolites [ 5 , 6 ] (metab-

olomics) provided unprecedented opportunities to unravel the

complexity of coordinated gene expression during conifer embryo

development. Analysis of the “omic” data now beneﬁ ts from the

extensive cDNA resources and proteome databases established in

several spruce and pine species of commercial and ecological inter-

est [

7 – 10 ]. It is also anticipated that the identiﬁ cation rate of mul-

tiple transcripts and proteins in genome-wide data sets will

considerably increase as decoding the conifer “giga-genome” is

ongoing in Picea glauca [

11 ], Picea abies [ 12 ], Pinus taeda [ 13 ],

and other pine species such as Pinus radiata (Scion-led project,

New Zealand, Scion Annual Report 2013, p. 15,

http://www.

scionresearch.com/__data/assets/pdf_file/0017/42443/

ScionAnnualReport2013-Highlights.pdf

), Pinus pinaster and

Pinus sylvestris (ProCoGen European project 2012–2015,

http://

www.procogen.eu/

). Even if annotation of this enormous genomic

resource is still challenging [

9 , 14 ], “omic” approaches to eluci-

date embryo development are rapidly developing. The resulting

knowledge might ultimately provide (epi)genomic tools for the

efﬁ cient production of improved and better adapted varieties

through seedlings and/or emblings ( somatic embryo genesis) to

cope with both market evolution and the changing environment.

It is also of particular interest to complement the tedious “trial and

error” strategy currently in use, to reﬁ ne somatic embryogenesis

protocols (reviewed by Klimaszewska et al., Chapter

7 ) and achieve

commercial application in conifer species, especially through mul-

tivarietal forestry [

15 , 16 ].

In this chapter we review the recent advances in transcrip-

tomics (and targeted gene expression studies), proteomics and

metabolomics of both somatic (SE) and zygotic embryo (ZE)

development in conifers. The review is mostly focused on SE, as

somatic embryo genesis has become a model in vitro system in

conifers to study the molecular biology of embryo development

[

2 , 17 ], including epigenetic aspects as background genetic load

can be ruled out from clonal material [

18 ]. It is comparatively dif-

ﬁ cult to sample manageable quantities of embryonal mass ( EM )

during early zygotic embryogenesis [

19 , 20 ]. Our aims were (1) to

emphasize the critical processes and genes at speciﬁ c embryo devel-

opmental and transition stages, and (2) to highlight practical appli-

cation for somatic embryogenesis in conifers.

2 Transcriptomics of Conifer Embryo Development

The development of cDNA or oligonucleotide-based microarray

technologies, and more recently, of next generation sequencing

RNA methods has provided critical advances for genome-wide

Jean-François Trontin et al.

169

screening of quantitative gene expression in forest trees [ 3 ].

Quantitative real-time polymerase chain reaction (qPCR) also pro-

vides an accurate tool for analyzing the expression of individual can-

didate genes selected from microarray and sequencing-based data.

The increasing use of absolute quantiﬁ cation may raise the possibil-

ity for universal comparison of gene expression [

21 ]. There are still

limitations at both technical [

22 ] and data interpretation levels [ 9 ].

However, microarrays developed in major conifers were found suit-

able for transcriptome proﬁ ling in other species [

9 , 23 ]. Both

microarrays and RNA sequencing methods are now increasingly

used to obtain either initial insights into embryogenesis- related

transcriptomes when public resources are limited, e.g., Larix

kaempferi [

24 ], or to perform comprehensive transcriptome analy-

ses of somatic [

2 , 18 , 25 ], and/or zygotic material [ 19 , 26 , 27 ].

Despite various spatiotemporal differences between gymnosperms

and angiosperms embryogenesis [

28 ], it is increasingly debated

how to efﬁ ciently translate the molecular information gained in A.

thaliana to domesticated species [

29 ]. The information delivered

from the completed genome sequence in A. thaliana resulted in a

deeper understanding of complex, regulated gene network (300–

450 genes) involved in embryo patterning [

20 , 29 – 31 ].

Interestingly, most embryogenesis-related genes identiﬁ ed in A.

thaliana have homologous sequences with strong congruity in

conifers [

28 ] such as in P. taeda (83 %) [ 32 ] and L. kaempferi (78

%) [

24 ]. Differences in molecular regulation of embryogenesis

between A. thaliana and conifers may therefore mainly arise from

variation in gene expression [

2 , 28 ], especially temporal differences

at the transition between embryo development al stages [

33 ].

Regulation of gene expression is thought to result partly from epi-

genetic modiﬁ cations as a possible adaptive mechanism in long-

lived trees [

18 ]. A large transcriptomic study further indicated that

ZE transcript proﬁ les are highly correlated between P. pinaster and

A. thaliana [

19 , 34 ], with gymnosperm -speciﬁ c transcripts esti-

mated to be only 3 %. There is thus some evidence that conifers

will beneﬁ t from angiosperms reference data [

20 , 35 , 36 ]. Because

transcripts abundance is not always predictive of effector molecules

underpinning the physiological process involved in embryo devel-

opment, one more challenge is to integrate transcriptomic with

proteomic and metabolomic datasets. This promising systems biol-

ogy approach to modeling the genetic regulation of plant embryo-

genesis is ongoing in A. thaliana [

20 ], with expected similarity in

conifers [

26 , 31 , 37 , 38 ].

Only a few available reviews [

28 , 39 – 43 ] aimed at unraveling the

complex regulatory gene network expressed in conifers, from

embryogenesis induction to early embryogenesis (embryonic

phase), late embryogenesis (up to the cotyledonary embryo stage)

2.1 Conifer

Transcriptomic

Analyses Beneﬁ t

from Advances

in Model Plants

2.2 The Growing

Bulk of Information

About Coordinated

Gene Expression

Molecular Aspects of Conifer Embryo Development

170

and the subsequent maturation steps from the acquisition of desic-

cation tolerance to the establishment of dormancy and accumula-

tion of storage reserves needed for germination . The range of

genes transcribed in conifer EM is apparently 30–40 % larger than

in any other tissue [

18 , 28 ]. Our knowledge is currently highly

fragmented because most studies performed during the last decade

were targeting few genes (Table

1 ). Little could be learnt until

recently about the expression of gene cohorts with similar tran-

script signatures during embryo development . For this purpose,

the comparative analysis of favorable and unfavorable SE matura-

tion conditions, as well as embryogenic and non-embryogenic

material, appeared critical to identify differentially expressed genes

among developmental stages [

1 , 21 , 23 , 46 , 89 , 90 ]. Most avail-

able transcriptomic data (Table

2 ) are from P. abies because SE

development is tightly controlled in this species with clear,

synchronized transition between developmental stages promoted

by speciﬁ c plant growth regulator ( PGR ) treatments. Transcript

proﬁ les were described from early embryogeny at the time of EM

proliferation to the cotyledonary stage using macroarrays [

23 , 90 ],

microarrays [

2 ], or sequencing methods [ 18 ]. The latter study spe-

ciﬁ cally reported on temperature-dependent differential transcrip-

tomes in proliferating EMs that may be associated with the

formation of an epigenetic memory with a delayed impact on seed-

ling development. In the closely related species P. glauca , Rutledge

et al. [

21 ] provided the ﬁ rst results of transcriptome analysis

(microarray) of early molecular events involved in the induction of

somatic embryo genesis in conifers. Stasolla et al. [

92 ] performed a

macroarray analysis of the effect of polyethylene glycol ( PEG ) dur-

ing SE maturation in P. glauca . Several macroarray studies also

investigated gene expression during early cotyledonary SE devel-

opment in P. radiata [

1 , 89 ], and from early embryogenesis to

cotyledonary SE and/or ZE development in P. taeda [

26 , 27 ]. In

P. pinaster , Morel et al. [

25 ] described differences in both gene

expression (sequencing) and proteome during early cotyledonary

SE development in favorable (high gellan gum concentration) and

unfavorable (low gellan gum) maturation conditions, whereas de

Vega-Bartol et al. [

19 ] performed the ﬁ rst microarray-based tran-

scriptomic proﬁ ling of ZE in a conifer, from early embryo to the

cotyledonary stage. Such transcriptomic studies excluded most

small RNAs for technical considerations. However, both microR-

NAs (miRNAs) and other small noncoding RNAs are part of the

epigenetic regulation complex of gene expression, which has a cru-

cial role in regulating development, including embryogenesis [

85 ,

106 ]. A high-throughput sequencing strategy was used in

L. kaempferi [

50 ] to identify miRNAs involved in regulation of

target genes at speciﬁ c SE stages. More than 100 predicted genes

were found to be putative targets of 60 miRNAs.

Jean-François Trontin et al.

171

Table 1

Recent targeted gene expression studies of embryo development in conifer species

Species Embryogenesis step Gene investigated Reference

Araucaria

angustifolia Early embryogenesis (SE) SERK 1 ( AaSERK1 ) [ 44 ]

Early to late embryogenesis (SE) Argonaute ( AaAGO ), CUC 1 ( AaCUC ), WOX ( AaWOX ), LEC1-like

( AaLEC ), vicilin 7S ( AaVIC ), S-locus lectin protein kinase ( AaLecK ),

reversible glycosylated polypeptide ( AaRGP ), scarecrow-like ( AaSCR )

[

45 ]

Larix

kaempferi Early embryogenesis (SE) miRNA ( miR159 , 169 , 171 , 172 ) [

46 ]

Early to late embryogenesis (SE) MYB-like ( MYB33 ), miR159 [

47 ]

Scarecrow-like ( SCL6 ), miR171 [

48 ]

Superoxide dismutase ( SOD ), catalase ( CAT ), ascorbate peroxidase ( APX ) [

49 ]

Trans-acting small interfering RNA TAS3/miR390 , dicer-like 1 DCL1/

miR162 , laccase/miR317 , plastocyanin/miR398 , ARF/miR160 , 167 ,

class III HD-ZIP/miR166 , miR156 , 159 , 168 , 171 , 397

[

50 ]

Larix x

marschlinsii Early to late embryogenesis, germination (SE) Germin-like protein 1 ( LmGER1 ) [

51 ]

Apetala 2-like ( LmAP2L1 , 2 ) [ 52 ]

Picea abies Early to late embryogenesis (SE) Type II plant metacaspase subfamily ( mcII-Pa ) [

53 ]

Actin isoformes ( Pa1 , 2 , 3 , 4 ) [

54 ]

KNOX1 ( HBK3 ), argonaute ( PgAGO ) [

55 ]

KNOX1 ( HBK1 , 2 , 3 , 4 ) [ 56 ]

Aquaglyceroporin ( PtNIP1;1 ) [

57 ]

HD-GL2 homeobox ( PaHB1 ) [

58 ]

CUC ( PaNAC01/02 ) [

59 ]

ABI3/viviparous 1 ( PaVP1 ) [

60 ]

ABI3/viviparous ( PaVP1 ), LEC1-like ( PaHAP3A ) [

61 ]

Early to late embryogenesis (SE/ZE) WOX ( PaWOX2 , PaWOX8/9 ) [

17 ]

Early to late embryogenesis, germination (SE) Lipid- transfer protein ( Pa18 ) [ 62 ]

Early to late embryogenesis (SE), germination (seeds) Class IV chitinase ( Chia4-Pa1 ) [

63 ]

Early to late embryogenesis (SE), vegetative tissue

(seedlings and/or older trees)

Lipid- transfer protein ( Pa18 ) [

64 ]

HD-GL2 homeobox ( PaHB2 ) [ 65 ]

WOX ( PaWOX2 , 3 , 4 , 5 , 8A , 8B , 8/9 , 13 ) [

66 ]

WOX ( PaWOX2 ) [

67 ]

PIN family of auxin efﬂ ux transporter ( PaPIN1 ) [ 68 ]

PIN1-like auxin transport protein ( PIN1-like ) [

69 ]

(continued)

Molecular Aspects of Conifer Embryo Development

172

Species Embryogenesis step Gene investigated Reference

Picea glauca Initiation and early embryogenesis (SE) Apetala 2-like ( AP2-L2 ), LEC1-like ( CHAP3A ), IAA2-like , babyboom

( SAP2C ), SERK1-like , KNOX ( SKN1 , 2 , 3 , 4 ), ABI3/viviparous ( VP1 ),

WOX ( WOX2 )

[ 70 ]

Early to late embryogenesis (SE) Argonaute ( PgAGO ) [

71 ]

HD-ZIP ( PgHZ1 ) [ 72 ]

SABATH methyltransferase ( PgIAMT1 ) [

73 ]

Late embryogenesis ACS ( ACS1 , 2 , 3 , 4 ) [

74 ]

Early to late embryogenesis, germination (SE) Wuschel homeobox ( WUS ), LEC1-like ( CHAP3A ) [

75 ]

Pinus contorta Early embryogenesis (SE), vegetative tissue (somatic

seedlings, mature tree) WOX ( PcWOX2 ), LEC ( PcHAP3A ) [ 76 ]

Pinus oocarpa Early to late embryogenesis (SE) 26S proteasome subunit S2 ( RPN1 ), HD-ZIP ( HD-Zip1 ), receptor

protein kinase clavata- like ( CLV ), LEA, legumin- and vicilin-like

proteins

[

77 ]

Pinus pinaster Early to late embryogenesis (SE/ZE) Glutamine synthetase ( GS1a , GS1b ) [

78 ]

Early to late embryogenesis (SE), adventitious

caulogenesis (ZE) Receptor protein kinase clavata 1-like ( PipsCLV1L ) [

79 ]

Early to late embryogenesis (ZE) Nonspeciﬁ c lipid- transfer protein ( PpAAI-LTSS1 ) [

80 ]

Early to late embryogenesis (ZE), germination

(seedlings) Rab-related small GTP-binding protein ( PpRab1 ) [

81 ]

Pinus pinea Early to late embryogenesis (SE), adventitious

caulogenesis (ZE) Receptor protein kinase clavata 1-like ( PipiCLV1L ) [

79 ]

Pinus radiata Early embryogenesis (SE/seedlings) Otubain-like cysteine protease ( PrOTUBAIN ) [

82 ]

Early to late embryogenesis (SE) GRAS family ( PrSHR , PrSCL1 , and 13 other GRAS genes) [

83 ]

Pinus sylvestris Early to late embryogenesis (SE/ZE) Glutamine synthetase ( GS1a , GS1b ) [ 78 ]

ACS ( PsACS1 , 2 ) [ 84 ]

LEC1-like ( PsHAP3A ), ABI3/viviparous ( VP1 ) [

61 ]

Table 1

(continued)

Jean-François Trontin et al.

173

Species Embryogenesis step Gene investigated Reference

Pinus taeda Early to late embryogenesis (SE) Class III HD-ZIP ( HB15L ), ARF ( ARF8L ), argonaute-like ( AGO9L ),

MYB factor ( MYB33 ), scarecrow ( SCRL ), apetala 2-like ( AP2L ),

miR159 , 166 , 167 , 171 , 172

[ 85 ]

Early to late embryogenesis (ZE) LEA ( LPZ202 , 216 , LSP094 ), dehydrin [

86 ]

Early to late embryogenesis (SE/ZE) 26S proteasome regulatory subunit S2 ( RPN1 ), HD-ZIP ( HD-Zip 1 ),

receptor protein kinase clavata-like ( CLV ), LEA, legumin- and vicilin-

like proteins

[

77 ]

Early to late embryogenesis (SE/ZE), vegetative tissue

(seedlings) Aquaglyceroporin ( PtNIP1;1 ) [

87 ]

Late embryogenesis (ZE) ABA responsive ( ABI3 , 4 , 5 ), root development ( woodenleg , short root ,

scarecrow , hobbit , bodenlos , monopteros ) [

88 ]

Late embryogenesis (SE/ZE) Starch synthase ( LPZ049 ), small HSP ( LPZ091 ), HSP70 ( LPZ270 ), LEA

( LPZ202 , 216 ), XETG-like ( LPZ060 ), cyclic phosphodiesterase

( LPZ016 ), 40S ribosomal protein ( LPZ206 )

[

86 ]

SE somatic embryo , ZE zygotic embryo , ABA abscisic acid, ABI ABA-induced, ACS 1-aminocyclopropane-1-carboxylic acid synthase, ARF auxin response factor, CUC cup-shaped

cotyledon, GTP guanosine triphosphate, HD-GL2 homeodomain- glabra2, HD-ZIP homeodomain-leucine zipper, HSP heat shock protein, IAA indole-3-acetic acid, KNOX knotted-

like homeobox, LEA late embryogenesis abundant protein, LEC leafy cotyledon, miRNA microRNA, SERK somatic embryogenesis receptor kinase , WOX wuschel-related homeobox,

XETG xyloglucan endo-transglycosylase

Molecular Aspects of Conifer Embryo Development

174

Table 2

Genome-wide molecular proﬁ ling of embryo development in conifer species

Species Embryogenesis step Method Reference

Transcriptomics

Araucaria

angustifolia Early to late embryogenesis (SE/ZE) NGS (Illumina) [

91 ]

Larix kaempferi Early embryogenesis (SE) NGS (454 sequencing) [

24 ]

Early to late embryogenesis (SE) NGS (Illumina), sRNA library [

50 ]

Picea abies Early embryogenesis (SE) cDNA array (373 cDNAs) [

90 ]

17 K cDNA microarray [

2 ]

NGS (Illumina) [

18 ]

Early to late embryogenesis (SE) 2 K cDNA array (2178 cDNAs) [

23 ]

Picea glauca Initiation and early embryogenesis

(SE) 32 K oligo-probe microarray [

21 ]

Early to late embryogenesis (SE) 2 K cDNA array (2178 ESTs) [

92 ]

Pinus pinaster Early embryogenesis (SE) NGS (Illumina) [

25 ]

Early to late embryogenesis (ZE) 25 K cDNA microarray [ 19 ]

Pinus radiata Early to late embryogenesis (SE) cDNA-AFLP [

1 ]

Screening of cDNA library [

89 ]

Pinus taeda Early to late embryogenesis (SE/ZE) cDNA array (326 cDNAs) [

26 ]

cDNA microarray (326 cDNA) [ 27 ]

Metabolomics

Picea abies Early to late embryogenesis (SE) GC/MS [

93 , 94 ]

Picea glauca Early and late embryogenesis (SE) NMR spectroscopy [

5 ]

Pinus taeda Early embryogenesis (SE) GC/MS [

6 ]

Proteomics

Araucaria

angustifolia Early embryogenesis (ZE) 2D-PAGE + LC-MS/MS [

95 ]

Early embryogenesis (SE) 2D-PAGE + MS [

96 ]

Early to late embryogenesis (ZE) 2D-PAGE + MS [

97 ]

Late embryogenesis, germination

(ZE) 2D-PAGE + LC-MS/MS [

98 ]

Cunninghamia

lanceolata Early to late embryogenesis (ZE) 2D-DIGE + LC-MS/MS [

99 ]

Cupressus

sempervirens Early to late embryogenesis (SE) 2D-PAGE [

100 ]

Larix x eurolepsis Early to late embryogenesis (SE) 2D-PAGE + LC-MS/MS [

101 ]

Late embryogenesis (SE) 2D-PAGE + LC-MS/MS [

102 ]

Larix

principis-

rupprechtii

Early embryogenesis (SE) 1D SDS-PAGE + iTRAQ protein

labeling + LC-MS/MS [

103 ]

Picea abies Late embryogenesis, germination

(SE) 2D-PAGE + GC-MS [

93 ]

Picea glauca Early to late embryogenesis (SE) 2D-PAGE + LC-MS/MS [

4 ]

Pinus

massoniana Early embryogenesis (ZE) 2D-DIGE + ESI-MS/MS [

104 ]

Pinus pinaster Early embryogenesis (SE) 2D-PAGE + LC-MS/MS [

25 ]

Late embryogenesis (SE/ZE) 2D-PAGE + LC-MS/MS [

105 ]

SE somatic embryo , ZE zygotic embryo , AFLP ampliﬁ ed fragment length polymorphism, cDNA complementary DNA,

DIGE difference gel electrophoresis, 2D-PAGE two-dimensional polyacrylamide gel electrophoresis , ESI-MS/MS elec-

trospray ionization coupled with tandem mass spectrometry , GC-MS gas chromatography coupled with MS, iTRAQ

isobaric tags for relative and absolute quantitation, LC-MS/MS liquid chromatography coupled with MS/MS, NGS

next generation sequencing, NMR nuclear magnetic resonance, SDS sodium dodecyl sulfate, sRNA small RNA

Jean-François Trontin et al.

175

In both conifers and angiosperms , little is known about gene

expression during the early stages of embryogenesis, which is rec-

ognized to be critical for subsequent embryo development [

1 , 18 ].

The embryonic phase is described as the dedifferentiation process

of mature, totipotent cells from competent explant (i.e., respond-

ing to stress or PGRs) to embryogenic cells (embryogenesis induc-

tion), giving rise to rapidly proliferating new early SE resulting in

the establishment of embryo-generating culture [

20 , 42 ]. In coni-

fers, the competent explants are restricted to ZE with limited prog-

ress from seedlings and from juvenile or adult trees (Klimaszewska

et al., Chapter 7). SE initiation may apparently proceed through

cell dedifferentiation within a competent explant. When cleavage

polyembryony occurs within seed (e.g., in Pinus ), “initiation” may

be merely the prolongation of this process in vitro [

42 ].

It is a particularly difﬁ cult task to identify genes underlying

somatic embryo genesis induction as the resulting initiated early

SEs may rapidly express additional confusing genes involved in

proliferation, maintenance of embryogenic potential [

20 , 21 , 48 ,

50 ], and early embryo development [ 24 ]. Recently, Elhiti et al.

[

20 ] reviewed the “omic” data available in plants and provided a

short list of 12 genes that are most likely to be involved in embryo-

genesis induction, from cell dedifferentiation ( ARF19/auxin

response factor19 , PRC1/ polycomb repressive complex 1 , RGP-1/

reverse glycosylating protein 1 , HSP17/heat shock protein 17 ), expres-

sion of totipotency ( SERK1/somatic embryogenesis receptor-like

kinase 1 , LEC1/leafy cotyledon 1 , GLB1/plant hemoglobin , WUS/

wuschel , a member of the WOX gene family, CLF/curly leaf ), and

commitment to embryogenesis ( CDKA/cyclin-dependent kinase A ,

PRZ1/ adaptor protein involved in CDK regulation, and STM/

shoot meristemless, a gene encoding KNOX1/ homeodomain pro-

tein of the KNOTTED1-like class). Homologous genes were found

in conifers but it is still unknown if they have similar expression

patterns and functions [

28 ]. Using the A. thaliana protein inter-

actome database, Elhiti et al. [

20 ] further identiﬁ ed 51 proteins

that may be functionally associated with the expression of these

12 genes.

Strikingly, there is currently no molecular study in conifers

dedicated to the early steps of somatic embryo genesis initiation

from juvenile explants (ZE). Only the identiﬁ cation of P. glauca

somatic trees which shoot buds have been responsive to initiation

treatment has provided a unique opportunity to gain insights into

the molecular aspects of embryogenesis induction in conifers [

21 ,

70 ]. In addition to conifer homologs of important genes for

embryogenesis induction discussed above, i.e., SERK1 , LEC1 ,

WOX2 , and SKN1 , 2 , 3 , 4 ( KNOTTED genes), Klimaszewska et al.

[

70 ] studied the expression of genes with a recognized function

during early embryogenesis, including AP2-L2 ( apetala ),

Auxin/ IAA2 ( indole-3-acetic acid-like 2 ), SAP2C ( babyboom ), and

2.3 Putative

Regulated Genes

During Embryogenesis

Induction

Molecular Aspects of Conifer Embryo Development

176

ABI3/VP1 ( viviparous ). After 3–6 days of induction, competent

bud explants were downregulated for AP2-L2 , SERK1 , and SKN1-

4 and upregulated for IAA2 and SAP2C . After initiation, most of

these genes were expressed ( SAP2C , SERK1 , SKN1 , 2 , and 4 ) or

upregulated in early SEs ( LEC1 , WOX2 , and VP1 ). SERK1 , LEC1 ,

and WOX gene s were similarly found expressed in early SE of other

conifers [

2 , 17 , 44 , 45 , 61 , 66 , 67 , 76 ]. WOX2 has been suggested

as a possible marker of effective initiation in conifers [

67 , 70 ].

Further transcriptomic comparison of responsive and nonrespon-

sive genotypes could be performed during somatic embryogenesis

induction in P. glauca [

21 ]. Surprisingly, only a few of the 12 can-

didate genes described by Elhiti et al. [

20 ] for cell dedifferentia-

tion, totipotency , and commitment to embryogenesis were found

to be regulated, i.e., several ARF genes (including ARF19 ), HSP17

and various CDK genes. It is suggested that effective SE initiation

requires not only the activation of embryogenesis-related genes,

but also a moderate activation of genes typical of adaptive stress

response of explant to induction treatment. Some of the most

highly expressed genes during SE induction in nonresponsive gen-

otype encoded proteins (apoplastic class III peroxidase, cell wall

invertase, serine protease inhibitors) possibly involved in biotic

defense activation. Interestingly, the jasmonic acid pathway

involved in biotic defense elicitation seems to be activated during

both cell dedifferentiation and totipotency acquisition in plants

[

20 ]. Gene activation in bud explants from the responsive geno-

type was comparatively lower in magnitude and/or only transient.

Among the most upregulated genes, such an expression pattern

was observed for a gene ( DHN1 ) encoding a conifer-speciﬁ c group

2 of late embryogenesis abundant proteins (group 2 LEA), in

accordance with a well-supported role for dehydrins in adaptation

to environmental stress. An apoplastic class III peroxidase gene was

also activated following a similar transient pattern, suggesting that,

in contrast to nonresponsive genotype, cellular redox homeostasis

was rapidly restored after the initial oxidative burst promoted by

the induction treatment. Peroxidases are also involved in various

physiological processes associated with cell dedifferentiation and

totipotency (e.g., auxin metabolism, cell wall modiﬁ cation). They

were shown to accumulate early during the induction phase of

somatic embryogenesis in Picea species [

107 ]. Two unknown

genes encoding proteins containing repetitive segments rich in

threonine– glutamine (QT-repeat) or proline (Proline-rich) were

also persistently upregulated in responsive genotypes. As these

genes appeared to be conifer-speciﬁ c, no general conclusion could

be drawn as to their putative role in somatic embryogenesis induc-

tion. Additional transcriptomic studies are therefore required to

increase our understanding of the basic mechanisms governing the

highly complex embryonic phase in conifers.

Jean-François Trontin et al.

177

As previously observed in A. thaliana , transcriptomic proﬁ ling of

both SE [

1 , 2 , 23 , 50 , 90 , 92 ], and/or ZE [ 19 , 26 , 27 ] in conifers

has revealed global characteristic changes in gene expression dur-

ing transition to successive developmental stages. In P. abies , com-

parative studies of normal and developmentally arrested

embryogenic lines [

23 , 90 ], revealed a transcriptional repressive

state during EM proliferation in the presence of PGRs, followed by

more active gene expression at the onset of embryo trans-

differentiation from EM, and again a repression state at the time of

embryo development . The number of differentially expressed

genes increased as embryos were developing [

2 ]. Most transcripts

(92 %) were unique, suggesting that different sets of genes are

regulated at the proliferation/early embryo development and

early/late embryo transitions. Important changes in gene expres-

sion between consecutive SE stages were similarly detected in P.

radiata [

1 ] and P. glauca [ 92 ]. Considering ZE, general variations

over multiple embryo stages were revealed in P. taeda [

26 , 27 ].

Both similarities and differences were observed between somatic

and zygotic patterns, suggesting that transcriptomics could be a

useful tool to check SE quality [

26 , 27 , 77 ]. In P. pinaster , de

Vega-Bartol et al. [

19 ] revealed a large set of differentially expressed

sequences from early to cotyledonary embryo stages. Functional

categories associated with these genes clustered into nine different

proﬁ les, each suggesting a high level of gene co-expression at the

same developmental stage. As in P. abies , there is an apparent gen-

eral trend during P. pinaster embryogenesis towards massive gene

regulation at the transitions from early to pre-cotyledonary

embryos and from cotyledonary to fully mature embryos. Such a

pattern may originate from both transcriptional (especially tran-

scription factor s, TFs) and posttranscriptional regulation through

various epigenetic mechanisms, including transposable element-

mediated DNA methylation and heterochromatin maintenance

(histone deacetylase genes) at early stages, large chromatin-

remodeling events during late embryo development, and ubiqui-

tous small RNA-mediated regulation (especially miRNAs) ac ross

all developmental stages.

Below we review major processes that have been highlighted in

transcriptomic studies for their crucial roles in the developmental

switch from early to mature embryo, including programmed cell

death ( PCD ), megagametophyte function and signaling, cell wall

modiﬁ cation, auxin signaling and other developmental regulator s,

abscisic acid ( ABA )-mediated processes, changes in metabolisms

(especially carbohydrates and proteins) and stress-related genes.

This is in close agreement with embryogenesis-related functions

supported by proteomic studies ( see Subheading

3 ).

As part of initial embryo polarization, suspensor cells and EM dif-

ferentiate very early in P. abies . Expression of a transmembrane

protein C gene ( TMP-C ) encoding an aquaglyceroporin, known to

2.4 Developmental

Switch

from Embryonic

to Vegetative Growth

2.4.1 Developmentally

Regulated PCD During

Somatic Embryogenesis

Molecular Aspects of Conifer Embryo Development

178

be predominantly localized in suspensor cells in both P. taeda [ 87 ]

and P. abies [

57 ], had already increased 24 h after early somatic

embryo s were stimulated to develop [

2 ] . TMP-C expression con-

tinued to increase with concomitant enlargement of suspensor size

up to the onset of exposure to ABA . Two waves of overlapping,

apoptotic and autophagic types of PCD are required for the appro-

priate development of SE, including degradation of proliferating

early SEs at the time of EM-to-SE transition and elimination of

terminally differentiated suspensor cells during early embryo mat-

uration [

108 – 110 ]. PCD is also activated during germination

[

108 , 111 ]. Upregulation of a cyclin-dependent kinase gene

( cdc2Pa ) involved in the progression of cell division was associated

with these periods of PCD. Kinase activity may initiate apoptosis

by phosphorylation of pro-apoptotic pr oteins [ 111 ]. Reorganization

of cytoskeletal structures also has an important role in PCD [

112 ],

and both actin and tubulin genes are regulated during conifer

embryogenesis [

23 – 25 ]. The actin cytoskeleton was reported to

differ between EM and suspensor cells with speciﬁ c expression in

suspensors of different actin isoforms [

54 ]. Application of low

doses of latrunculin B (an actin depolymerizing drug) during SE

maturation predominantly degraded suspensor cells, which in turn

accelerated and synchronized the development of high-quality

embryos [

54 ]. Actin depolymerisation has been shown to induce

PCD associated with caspase-like activities in plants [

113 ]. A gene

encoding a putative actin depolymerizing factor is upregulated in

P. abies at the early stage of embryo development [

23 ]. PCD was

found to be activated as soon as PCD-related genes encoding tran-

sient VEIDase/caspase-like activity, such as cathepsin B-like cyste-

ine protease or metacaspase , were signiﬁ cantly upregulated at the

EM-to-SE transition. Aquea and Arce-Johnson [

1 ] similarly found

an uridylate kinase gene and a type-II metascaspase gene upregu-

lated during early embryo development in P. radiata . Metacaspase

genes are recognized candidates for performing the role of cysteine

protease genes [

53 ], whereas speciﬁ c alterations in the balance of

pyrimidine nucleotide synthesis, involving uridylate kinase, may

represent an early signal for PCD [

1 ]. Endochitinase genes have

also been associated with PCD in plants [

63 ] and were found to be

upregulated in L. kaempferi embryogenic cultures [

24 ], during

embryo development in P. abies ( Chia4-Pa ) [

63 ], and at the onset

of embryo development in P. pinaster [

25 ]. Chia4-Pa expression

was found to be restricted to the EM (embryo proper) base, and

chitinase accumulated to form a covering ﬁ lm on the whole EM

surface. As observed in Pinus caribaea [

114 ], chitinases apparently

target speciﬁ c arabinogalactan protein s located in the epidermal

cell wall [

63 ]. In maturing EMs of P. pinaster , regulation of PCD

was also suggested by the increased expression of a disulﬁ de isom-

erase gene encoding a protein known to interact with speciﬁ c cys-

teine proteases [

25 ].

Jean-François Trontin et al.

179

Large suspensors are formed by conifer embryos [ 26 ]. Although it

remains a transient organ, the suspensor is central to embryo devel-

opment , which includes physical support, and the translocation

and synthesis of nutrients and signaling molecules. Transcriptomic

analysis of pre-cotyledonary ZEs in P. taeda revealed that various

genes, encoding proteins normally associated with late embryo

development (e.g., storage proteins , LEAs), are upregulated in

suspensor tissue when compared to EM . This expression pattern

has signiﬁ cant similarities with that of the megagametophyte , sug-

gesting that the suspensor may be involved in the production of

storage and other compounds to be mobilized during embryo

development [

26 ]. Various homologous genes of putative signal-

ing factors, normally expressed in the female gametophyte, were

found to be upregulated ( ATHB22/MEE68 , MEE49 ) or down-

regulated ( MEE66 ) from early-embryo to late-embryo develop-

ment in P. abies [

2 ]. These genes are known to affect both

endosperm and early embryo patterning in A. thaliana , suggesting

that some somatic cells in proliferating EMs may have a megaga-

metophyte signaling function in conifers. Endosperm properties

are similarly recognized for both class IV chitinase ( Chia4-Pa )

[

63 ] and NARS2 [ 2 ] genes upregulated during early embryogen-

esis ( see Subheadings

2.4.1 and 2.4.4 ), as well as for genes highly

expressed at the onset of late embryo development in P. pinaster

and involved in posttranscriptional regulation of gene transcription

(small ubiquitin-related modiﬁ er/SUMO- or ubiquitin-

conjugating enzyme) [

25 ]. From in situ observations indicating

that Chia4-Pa genes are expressed in subpopulations of cells in

both proliferating EMs and early embryos [

63 ], it is speculated

that “nurse” cells expressing developmental regulator s with mega-

gametophyte signaling functions are required in conifers [

2 , 63 ].

The auxin-mediated effect on reorganization of cell wall architec-

ture, possibly through alterations in cytoskeleton structure, is well

established and has implications in cell fate and differentiation

[

23 ]. Many developmental regulator s involved in embryo pattern-

ing proceed through cell wall modiﬁ cations. Most transcriptomic

studies have revealed that these modiﬁ cations are developmentally

regulated from embryogenesis induction to early and late embryo-

genesis, thus supporting their role in proper embryo development .

In P. abies , several genes encoding enzymes involved in the synthe-

sis of hemicellulose and pectin (UDP-glucose dehydrogenase), low

molecular weight galactosides and cell wall polymers (UDP-

galactose 4-epimerase like) were downregulated, especially at the

time of effective early embryo development. A laccase gene involved

in ligniﬁ cation and thickening of the cell wall was also downregu-

lated during the transition from pre-cotyledonary to cotyledonary

embryo [

23 ]. A similar expression pattern was observed in L.

kaempferi [

50 ] and correlated with expression of miR397 ,

2.4.2 Genes

with Megagametophyte

Function and/or Signaling

2.4.3 Genes Related

to Cell Wall Modiﬁ cation

Molecular Aspects of Conifer Embryo Development

180

supporting a posttranscriptional regulation of laccase during

somatic embryo genesis. Additional genes involved in cell wall loos-

ening and reorganization are regulated at early stages in P. radiata

[

1 ] and P. abies [ 2 ], such as genes encoding α- d -galactosidase and

myo-inositol oxygenase, expansin, and pectinesterase. Moreover, a

β-expansin gene was found speciﬁ cally expressed in EM [

89 ], as

well as genes encoding cellulase and apoplastic germin-like protein

(GLP). At the switch from early- to late-embryo stages in P. abies ,

different expansin and pectinesterase genes, as well as a

xyloglucan:xyloglucosyl transferase gene, signiﬁ cantly changed their

expression level [

2 ]. Transcriptomic and proteomic analyses at the

onset of late embryogenesis in P. pinaster [

25 ] similarly revealed

overexpression of cell wall-related expansin genes and also a drastic

upregulation of a putative gene encoding extensin-like protein.

Auxin biosynthesis and relocalization by polar auxin transport has

a crucial function in the activation of the auxin response machinery

during plant embryogenesis that results in setting up (1) apical–

basal patterning (meristematic poles) and (2) radial embryo pat-

terning (adaxial/abaxial organization). Various reports suggested

that auxin-mediated events are of similar high importance in both

angiosperms and gymnosperms to establish a roughly similar basic

body organization. A recent transcriptome comparison of early SE

and ZE performed in A. angustifolia suggested that incomplete SE

development resulted from an auxin signaling failure in embryo-

genic cultures [

91 ]. Endogenous auxin biosynthesis (especially

IAA) is activated early during SE development in P. abies [

69 ],

whereas putative auxin transport proteins are upregulated [

23 ].

Concomitantly, auxin starts to be relocalized by polar transport as

observed in both P. abies [

115 ] and P. sylvestris [ 116 ]. Disruption

of polar auxin transport by N -1-naphthylphthalamic acid (NPA)

affected early embryo polarization, promoted aberrant develop-

ment, such as no or poor shoot apical meristem (SAM) and fused/

aborted cotyledons, and resulted in abnormal germination [

69 ,

115 , 116 ]. Upregulation of auxin-responsive gene ( SAUR ) and

downregulation of auxin biosynthesis competition gene ( SUR1 ,

involved in glucosinolate synthesis, a sister branch of IAA biosyn-

thesis) were indicative of increased auxin synthesis during develop-

ment of early stage embryos in P. abies [

2 ]. IAA homeostasis may

be modulated by additional mechanisms such as methylation of the

free carboxyl group by methyltransferases of the plant SABATH

family. Such a gene showing high catabolic activity with IAA was

expressed in P. glauca during early embryo development , and then

downregulated towards later stages [

73 ]. Expression of LEC1 dur-

ing early embryogenesis in P. abies and P. sylvestris , followed by a

strong decrease at the switch to late embryogenesis, was also

observed [

2 , 61 ]. The activation of genes involved in localized

auxin biosynthesis has been linked with expression of both LEC1

2.4.4 Auxin Response

Machinery and Embryo

Patterning

Jean-François Trontin et al.

181

and LEC2 [ 20 , 75 ]. In P. pinaster ZE, a TF gene orthologue to the

auxin response factor ( ARF16 ) involved in regulation of auxin-

modulated genes (e.g., WOX5 : maintenance of pluripotent cells in

root quiescent center) was drastically downregulated at the transi-

tion to pre-cotyledonary embryos [

19 ]. In L. kaempferi , expres-

sion of different ARF genes, possibly regulated by miR160 or

miR167 , increased up to the early SE cotyledonary stage and then

signiﬁ cantly decreased [

50 ]. In the same species a putative trans-

acting small interfering RNA (siRNA) gene ( TAS3 ), regulated by

miR390 and known to target several ARF genes, was differentially

expressed from early to late embryogenesis. TAS3 has been involved

in the juvenile-to-adult phase transition through the negative reg-

ulation of ARF genes [

117 ]. Zhang et al. [ 50 ] similarly observed

the upregulation at the cotyledonary stage of miR156 , targeting a

crucial gene for the juvenile-to-adult transition ( SPL3 ). It is

strongly suggested that auxin-mediated, early cell fate decisions,

such as root apical meristem (RAM) delineation, are contributing

to apical-basal embryo polarization. During the same transition,

upregulation of a putative TF gene from the KANADI family

( KAN2 ) involved in the regulation of polar expression of auxin

efﬂ ux-facilitating proteins genes from the PIN-FORMED family

(PIN), as well as concomitant regulation of PIN3 and a gene

involved in the recycling of PIN proteins ( GNOM ), indicated

active modulation of auxin ﬂ ow. Interaction of KAN genes and

class III HD-Zip (class III homeodomain leucin zipper) TFs with

auxin has been involved in abaxial pattern formation, especially

during emergence of cotyledon primordia. Accordingly, a HD-Zip

III gene target of miR166 was found to be upregulated at the early

cotyledonary stage in L. kaempferi [

24 ]. Furthermore, TF genes

known as primary coordinators of polar auxin transport (auxin

inﬂ ux carrier, AUX1 ) and modulation of auxin transport ( NDL1 ),

possibly through regulation of AUX1 and other PIN protein genes

( PIN2 ), were also upregulated at early stages [

19 ]. If such impor-

tant TFs have conserved functions in plants, they may contribute

to auxin-related spatiotemporal regulation of genes that are

involved in the establishment of early embryo patterning, as well as

activation of the auxin response machinery later during develop-

ment. In conifers, most TF genes are probably regulated by miRNA

themselves [

24 ].

Expression of auxin-induced genes signiﬁ cantly increased

from early cotyledonary to cotyledonary embryos in P. glauca

[

92 ], as was ARF16 in P. pinaster [ 19 ]. Vestman et al. [ 2 ] found

that the putative conifer SUR1 is upregulated at the onset of late

embryo development , while expression of SAUR was maintained

at high levels, suggesting an increase in glucosinolate biosynthe-

sis together with maintenance of a high IAA level. Upregulation

of genes encoding the auxin-induced protein (IAA11), auxin

receptor (TIR1), TF regulator of auxin-responsive gene

Molecular Aspects of Conifer Embryo Development

182

(MYB77) and a positive regulator of brassinosteroid signaling

suggested that auxin-responsive gene expression is being acti-

vated. At the transition from pre-cotyledonary to early cotyle-

donary embryos in P. pinaster ZE, a putative TF gene regulated

by auxin and involved in SAM function ( ANT ) was signiﬁ cantly

overexpressed. At later stages, upregulation of a putative TF

gene from the YABBY family ( YABBY2 ) with polar expression

resulting from interplay with KAN and phabulosa genes is con-

sistent with determination of adaxial–abaxial cell fate [

19 ].

Similarly, downregulation in cotyledonary embryos of a putative

TF gene, required for the establishment of leaf primordia adax-

ial–abaxial polarity ( AS2/LOB ) and repression of meristem-

related homeobox genes of the KNOTTED1-like class ( KNOX1 ),

indicated that the formation of SAM and organ boundaries had

started. Expression of some KNOX1 genes in P. abies ( HBK2 ,

HBK4 ) is speciﬁ c of competent EM to form cotyledonary

embryos [

56 ]. Delayed expression of HBK2 and HBK4 in NPA-

treated lines resulted in embryos lacking SAM. Similarly,

KNOTTED-like genes upregulated during early embryogenesis

are downregulated at later stages in P. glauca [

92 ].

Upregulation of a putative member of the NAC domain (spe-

ciﬁ c DNA-binding domain in the N-terminal region) TF family,

which is involved in downstream auxin signaling ( NAM/NARS2 ),

further suggested that SAM formation was initiated. NARS2 is

also upregulated in P. abies from early embryo development to the

onset of late embryogenesis [

2 ], indicating that delineation of

important tissue might be ongoing as embryos start to develop.

NAM and also other members of the large NAC domain TF family

regulated by PIN1 (auxin carrier proteins), such as CUC (cup-

shaped cotyledon) genes 1 and 2, are crucial for the initiation of

SAM, as well as the formation and separation of aerial organs. A

member of the polycomb group (Pc-G) protein (curly leaf, CLF ),

a part of the polycomb repressive complex 2 (PRC2) involved in

chromatin remodeling , was increasingly upregulated towards the

mature ZE stage in P. pinaster [

19 ]. Both CUC2 and PIN1 genes

have been described as target genes for Pc-G proteins. Expression

of a NAC homologue of CUC1 and CUC2 in P. abies ( PaNAC01 )

was regulated by polar auxin transport and was associated with

SAM differentiation and formation of separated cotyledons [

59 ]. A

CUC1 -like gene was similarly regulated during maturation of SE

in Araucaria angustifolia [

45 ], as was a PIN1 -like gene from pre-

cotyledonary to cotyledonary SE stages in P. abies [

68 , 69 ]. The

embryo apical parts accumulated more IAA, especially in the pro-

todermal cell layer where PIN1 -like expression was high. NPA

treatment of embryos before cotyledon initiation disrupted this

pattern and resulted in deregulation of both PIN1 -like [

69 ] and

WOX2 [

118 ], one of the WUS/WOX TF family members acti-

vated during embryo development [

17 , 67 ]. It is suggested that

Jean-François Trontin et al.

183

correct auxin transport is crucial at the transition from early to pre-

cotyledonary embryos and is involved in the coordinated regula-

tion of WOX2 and PIN1 . Polar auxin transport may proceed

through actin-dependent PIN proteins cycling between cytoplas-

mic membrane and the endosomal compartment. A putative gene

encoding a small Rab-related GTP-binding protein ( PpRab1 )

involved in ER-to-Golgi vesicle transport was differentially

expressed throughout embryo development in P. pinaster [

81 ].

Differential expression of WOX members, as a function of auxin

ﬂ ow and through a regulatory loop with CLAVATA1 ( CLV1 ), has

been proposed as one mechanism contributing to delineation of

different embryo domains [

2 , 17 , 68 ]. CLV1-like genes are

apparently expressed from early to late SE development in P. glauca

[

92 ], and P. pinaster and Pinus pinea [ 79 ].

Several genes known as important developmental regulator s dur-

ing early somatic embryo genesis, such as SERK1 ( cell reprogram-

ming ) and WOX2 ( cell fate decision, domain delineation), were

found continually expressed during SE development in P. abies [

2 ].

A similar pattern was observed with genes involved in the organi-

zation of cell division ( FK / fackel , RBR1/retinoblastoma-related1 )

and SAM formation ( PNH / pinhead ), suggesting that embryo pat-

terning is to some extent organized from the early embryo stages.

PNH is a member of the Piwi Argonaute ZWILLE family known

to act together with AGO1 , a member of the argonaute family tak-

ing part in the RNA-induced silencing complex. In conifers, AGO

gene s are required for embryo development [

19 , 71 , 85 ] and are

themselves regulated by miRNA (e.g., miR168 ) [

50 ]. In P. pinas-

ter ZE, AGO genes were highly represented in late embryos and

may be mediators of either 24-nt-long siRNA ( AGO9 , silencing of

transposable/repetitive elements) or miRNA and other siRNAs

( AGO1 ). Several other genes were regulated towards late stages,

such as upregulated dawdle ( DDL ) and hyponastic leaves1 ( HYL )

or downregulated dicer-like1 ( DCL1 ) and ﬂ owering locus CA

( FCA ) [

19 ]. A decrease of DCL1 expression was similarly observed

in L. kaempferi with feedback regulation by miR162 [

50 ]. High

expression of DCL1 at early stages may prevent precocious expres-

sion of important TFs through TAS-derived, siRNA-triggered

DNA methylation [

19 ]. AGO-like genes were upregulated at early

embryo stages in both P. glauca [

92 ] and A. angustifolia [ 45 ]. In

P. glauca AGO genes were preferentially expressed in SAM and

RAM and deregulation resulted in severe embryo abnormalities

[

71 ]. Overexpression of a KNOX1 gene ( HBK3 ) in P. abies prolif-

erating EMs resulted in the upregulation of AGO and accelerated

SE development with enlarged SAM areas [

55 ]. Proper SAM for-

mation in P. glauca SE was also suggested by overexpression of

related ZWILLE (stem cell maintenance within SAM along with

WUS/WOX ), KNOTTED -like ( see Subheading

2.4.4 ) and

2.4.5 Other Important

Regulators for Early

Establishment of Embryo

Body Plan

Molecular Aspects of Conifer Embryo Development

184

FIDDLEHEAD genes ( FDH , cell division and differentiation).

Developmental regulator s differentially expressed in P. abies [

2 ],

and supporting that embryo pattern formation starts very early,

include homologues of PDF2 and LUG , LBD12/ASL5 , LBD15 ,

and LBD40 . Expression of a PDF2-like gene (protodermal factor 2,

an HD-GL2 homeobox gene) in P. abies protodermal cells ( PaHB1

gene) is an indicator that SE protoderm will be formed [

58 ]. The

expression pattern of a lipid transfer protein (LTP) gene ( Pa18 )

was also associated with differentiation of protoderm and adjacent

outer cell layers [

64 ]. A similar nonspeciﬁ c LTP gene ( PpAAI-

LTSS1 ) was upregulated from pre- to early-cotyledonary ZE in P.

pinaster [

80 ]. Deregulation of Pa18 in proliferating EM negatively

impacted SE morphology and growth [

62 ]. A different HD-GL2

gene ( PaHB2 ) is also expressed in proliferating EMs and early SE

but its pattern becomes restricted to the subepidermal cell layer in

the mature embryo [

65 ]. This gene could be involved in speciﬁ ca-

tion and maintenance of the cortex identity. Both PaHB1 and

PaHB2 are suggested markers to monitor radial pattern formation

in P. abies . Another important gene in early delineation of radial

patterning (embryonic root) is SCR from the GRAS TF family

( scarecrow ). SCR expression was upregulated in P. glauca EMs

shortly after transfer to maturation conditions [

92 ], as well as in

cotyledonary embryos from P. taeda (ZE) [

88 ] and P. radiata

(SE) [

83 ]. SCR-like members appeared to be regulated by miR171

in L. kaempferi [

46 , 48 , 50 ]. Genes encoding LOB domain-

containing (LBD) and/or asymmetric leaves-like (ASL) proteins

(LBD12/ASL5, LBD15, LBD40) are identiﬁ ed as regulators of

the formation of lateral shoot boundary regions ([

2 ], and refer-

ences therein). LUG may have an early role in the speciﬁ cation of

EM cells by contributing to prevent ectopic expression of homeo-

tic AGAMOUS gene. ROXY1 is another negative regulator of

AGAMOUS , downregulated at the switch from the early to late

embryo stages [

2 ]. Expression of LEC gene s can directly induce

AGAMOUS during early embryogenesis, which in turn upregu-

lates gibberellin 2-oxidase ( GA2OX gene) and decreases gibberel-

lic acid (GA) synthesis [

20 ]. Downregulation of LEC1-like genes

was observed at the transition from early to late embryo [

2 ].

Increased GA biosynthesis was concomitantly supported by down-

regulation of GA2OX , one gene involved in GA catabolic process

and upregulation of negative regulators of GA signaling ( SPINDLY ,

cotyledon formation) and response pathways ( RGL1 ). As LEC

genes are also associated with auxin biosynthesis [

20 , 75 ], it

became apparent that expression of LEC and other regulators of

AGAMOUS are involved in spatiotemporal modulation of auxin-

and GA-mediated responses. The temporal and organ-speciﬁ c

expression of homeotic genes such as the coordinated AGAMOUS

and APETALA2 [

52 ] therefore appeared to have direct implica-

tion in embryo patterning. Both genes are likely to be controlled

Jean-François Trontin et al.

185

by epigenetic regulator s, such as MSI1 encoding a core protein of

the PRC2 complex similarly regulated in A. thaliana and P. pinas-

ter [

19 ]. Part of the complex machinery involved in embryo pat-

terning is therefore conserved between angiosperms and

gymnosperms [

2 , 19 ].

Global alteration in gene expression was observed in EM matured

on ABA -containing medium in conifers [

1 , 25 , 27 , 92 ]. In combi-

nation with other triggers (e.g., sucrose , PEG , gellan gum), ABA

stimulates the development of late stage embryos. ABA may alter

EM responsiveness to PGRs (auxin, GA) and may promote estab-

lishment of the embryo body plan [

92 ]. An exogenous supply of

ABA is needed in vitro as it is essentially provided by the megaga-

metophyte [

77 ].

As previously discussed, the key TF developmental regulator-

LEC1 is expressed during early embryogenesis [

2 , 45 , 61 , 70 , 75 ,

76 ], and then becomes signiﬁ cantly downregulated at the onset of

late embryo development promoted by exogenous ABA [

2 , 61 ].

The resulting putative modulation of both auxin- and GA-mediated

signaling pathways could be involved in the developmental switch

from embryonic to vegetative growth. LEC1 (HAP3 subunit) and

LEC2 (B3-domain) TF genes are part of a complex regulatory net-

work with additional B3-domain genes, such as ABI3 (ABA insen-

sitive 3) and FUSCA/FUS3 (fused cotyledon 3), resulting in direct

or indirect ABA-dependent regulation of genes [

2 , 28 , 119 ]. These

four genes are known as the master regulators of late embryogen-

esis in A. thaliana [

120 ]. They act synergistically to regulate the

expression of important downstream pathways, e.g., carbohydrate

metabolism, biosynthesis of storage proteins , LEAs or fatty acids.

FUS3 is a central regulator promoting increased endogenous ABA

synthesis while decreasing GA levels. The ABA signal transduction

cascade involved inactivation of ABA-insensitive (ABI) protein

phosphatases 2C (PP2C), promoting phosphorylation of serine/

threonine residues and activation of Sucrose non-fermenting 1

(Snf1)-related protein kinases 2 (SnRK2) and other calcium-

dependent kinases. SnRK2 subsequently activates downstream tar-

gets, especially ABA-response elements binding the HD leucine

zipper (B-ZIP) TF from the ABF/AREB/ABI5 clade [

120 , 121 ].

Expression of ABI3 and FUS3 is further triggered by exogenous

sugar [

120 ]. A FUSCA/FUS3 homologue was differentially

expressed in P. glauca during late SE development [

92 ]. Several

PP2C and SnRK2 transcripts were signiﬁ cantly expressed in P. pin-

aster and endogenous ABA level increased after 4 weeks of matura-

tion, suggesting an ontogenetic signal for SE development [

25 ].

ABI3 was upregulated at the transition to late SE development in

P. abies [

2 ] as well as ABI4 (sugar signaling), a B-ZIP TF, several

genes encoding LEAs, and a heat shock TFs. Homologues of

ABI3 , 4 , and 5 were similarly regulated in P. taeda [

88 ].

2.4.6 ABA -Mediated

Developmental Switch

from Embryonic

to Vegetative Growth

Molecular Aspects of Conifer Embryo Development

186

Upregulation of LEA and HSP genes was also observed at late SE

stages in P. abies [

23 ] and P. glauca [ 92 ] during SE and ZE devel-

opment in P. taeda and during SE maturation in Pinus oocarpa

[

77 , 86 ]. Both LEAs and HSPs have been associated with the

acquisition of embryo desiccation tolerance [

92 ] ( see Subheading

3 ). The fatty acid elongase gene FDH (epidermal cell differentia-

tion) and an extracellular dermal glycoprotein ( EDGP ) were also

downregulated [

2 ], as was a B-ZIP gene at the transition to early

ZE in P. pinaster [

19 ]. Similarly, one HD-ZIP I gene gradually

decreased with maturation of late SEs and ZEs in P. taeda [

77 ] and

P. glauca [

72 ]. In contrast, a lower but sustained HD-ZIP I expres-

sion was observed in maturing P. oocarpa SEs [

77 ]. Upregulation

of an ABI3 homologue ( PaVP1 ) was similarly reported in P. abies

and P. sylvestris [

60 , 61 , 111 ]. In P. abies , PaVP1 expression was

speciﬁ c to EM and was maintained at a high level in productive

lines until the cotyledonary stage [

60 ]. Both LEC1 and PaVP1

expression are affected by the histone deacetylase inhibitor tricho-

statin A, suggesting possible control by chromatin remodeling

[

61 ]. Several histone deacetylase genes were revealed as important

epigenetic regulator s regulated from early ( HD2C , HDA2 ) to late

embryogenesis ( HDA8/9 ) in P. pinaster [

19 ]. Other differentially

expressed genes involved in H3K9 methylation ( SUVH1 ) or

encoding chromatin-remodeling ATPases may have a similar role

in the organization of transcriptionally repressive chromatin. In P.

abies , Vestman et al. [

2 ] also revealed the downregulation of a gene

from the WRKY TF family, known to act downstream of the PP2C-

ABA receptor complex and to target ABA-responsive genes (e.g.,

ABF2-4 , ABI4-5 , MYB2 , DREB1a-2a , RAB18 ) [ 122 ]. Accordingly,

several genes activated by ABA are upregulated in P. abies [

2 ],

including putative DREB TFs (response to dehydration),

Angustifolia 3 ( AN3 ) and its target growth regulating factor 1

( GRF1 ), both involved in initial leaf morphogenesis, and a NAC

domain-containing protein gene ( ANAC009 ) expressed in grow-

ing tissues. Expression of the TF gene encoding MYB33 homo-

logue was shown to decrease during late embryo development

[

47 ]. MYB33 and other GAMYB-like genes are positive regulators

of ABA response and are essential for the temporal regulation of

development. Interestingly, MYB33 was found to be regulated by

miR159 in L. kaempferi as in A. thaliana . Expression of this

miRNA is induced by ABA in an ABI3-dependent mode ([

47 ],

and references therein).

During the transition from early to late SE development in P. abies ,

Vestman et al. [

2 ] observed an increase in carbohydrate metabolism

as revealed by the upregulation of phosphofructokinase 2 ( PFK2 )

involved in glycolysis and sucrose synthase 3 ( SuSy3 ) genes. In P.

glauca , many genes involved in sucrose catabolism (glycolytic and

tricarboxylic acid pathways) were downregulated at the SE

2.4.7 Changes

in Metabolisms

and the Link

with Important Processes

Jean-François Trontin et al.

187

cotyledonary stage [ 92 ]. Conversely SuSy was upregulated from

pre- to early-cotyledonary embryos. Activation of the glycolytic

pathway in unfavorable conditions for P. pinaster SE maturation

was revealed by transcriptomic, proteomic, and carbohydrate analy-

ses [

25 ]. Active carbohydrate catabolism at the onset of SE matura-

tion may preclude embryo development . In favorable maturation

conditions, various ubiquitine protein ligases genes were overex-

pressed, as well as ubiquitin-/small ubiquitin-related modiﬁ er

(SUMO)-conjugating genes [

25 ]. Ubiquitine protein ligases are

associated with SUMO activation (a chromatin modiﬁ er), suggest-

ing that maturing early SEs were subjected to global modiﬁ cations

of gene expression. Ubiquitin protein ligases are also activators of

the PGR -regulated ubiquitin/26S proteasome pathway resulting in

controlled proteolysis with increased supply of amino acids.

Accordingly, upregulation of a 26S proteasome subunit gene

( RPN1 ) was observed early in P. taeda [

77 ], suggesting that con-

trolled proteolysis is active at the developmental switch from early

to late embryo. This pathway is also activated in P. radiata early SE

and later embryo stages, as revealed by the expression of a gene

from the OTUBAIN family of cysteine proteases involved in remov-

ing the ubiquitin chain of protein destined for degradation [

82 ].

Transcripts, as well as corresponding storage proteins of the

legumin- and vicilin-like classes, were shown to increase with a

similar pattern in developing SE and ZE in P. taeda [

77 , 123 ] ( see

also Subheading

3 ). Interestingly, suboptimal conditions for SE

maturation in P. oocarpa and A. angustifolia resulted in lower

expression of genes encoding legumin- and/or vicilin-like storage

proteins [

45 , 77 ]. In P. glauca , deposition of storage proteins was

increased in cotyledonary SEs obtained on PEG -containing matu-

ration medium [

92 ], in accordance with the upregulation of two

genes encoding glutamine or glutamate synthase (GS/GOGAT

cycle). Activation of nitrogen assimilation through the GS/

GOGAT cycle may result in increased available glutamine for stor-

age proteins synthesis. Expression of the two glutamine synthase

genes, associated with green ( GS1a ) or vascular tissues ( GS1b ), was

studied in P. pinaster and P. sylvestris embryos. GS1a was expressed

in SE but not in ZE cotyledons, suggesting precocious SE germi-

nation [

78 ]. Expression of GS1b was detected in proliferating early

SE and in procambial cells of both cotyledonary SE and ZE, sug-

gesting that glutamine biosynthesis is effective a long time before

differentiation of mature vascular elements. GS1b expression was

proposed as a marker of SE quality [

78 ]. An increase in endoge-

nous polyamines levels (spermine, spermidine), another important

class of nitrogen compounds, was observed during SE maturation

in P. glauca ([

92 ], and references therein). Polyamines synthesis

requires decarboxyl-SAMet ( S -adenosyl-methionine). SAMet orig-

inates from methionine (SAMet synthase activity) and is also a

direct precursor of ethylene (ACC synthase/oxidase activities) and

Molecular Aspects of Conifer Embryo Development

188

a methyl donor in transmethylation mechanisms resulting in DNA

or histone methylation. Methyl residue transfer from SAMet results

in the production of S-adenosylhomocysteine (SAH), which is

recycled into methionine through production of adenosine (SAH

hydrolase). AMP is further produced from adenosine by adenosine

kinase (AK). Stasolla et al. [

92 ] found that an ACC oxidase gene

was upregulated at the SE cotyledonary stage in P. glauca whereas

several AK genes were downregulated, indicating active ethylene

synthesis. High expression of ACC synthase genes was also observed

in this species ( PgACS1 ) [

74 ] and in P. sylvestris ( PsACS2 ) [ 84 ],

especially at the early cotyledonary stage. PsACS2 expression was

proposed as a marker of competent embryo development in P. syl-

vestris since it was positively correlated with both ethylene produc-

tion and embryogenic potential . Vestman et al. [

2 ] similarly

reported a strong increase in ACC oxidase expression in P. abies at

the developmental switch to late embryogenesis. Upregulation of

SAH hydrolase and methionine synthase and expression of SAMet

synthase genes were also observed after induction of embryo devel-

opment [

23 , 90 ]. Expression of these genes supported active trans-

methylation events resulting in DNA or histone methylation that

may contribute to the global transcriptional repression state

observed at speciﬁ c embryo stages [

23 ]. In P. pinaster , both DNA

methylation and heterochromatin maintenance were important

processes at the onset of embryo maturation through transposable

element-speciﬁ c DNA methylation resulting in heterochromatin

formation ( DDM1 ), RNA-directed methylation of various trans-

posable elements ( FCA ), and regulation of DNA methylation

involved in regulation of chromatin structure ( ORTH2/VIM1 )

[

19 ]. In accordance with the increased level of SAMet synthase

proteins from early to late embryo stages in conifers ( see Subheading

3 ), normal embryo development appeared associated with the

modulation of polyamine and ethylene synthesis, as well as with

epigenetic regulation of gene expression.

As also revealed from proteomic and metabolomic studies ( see

Subheadings

3 and 4 , respectively), a general trend towards regula-

tion of genes involved in response to stress was observed at the

transition to early embryo development and also during late devel-

opment. The modulation of gene response to stress is required for

proper embryo development and could be of practical interest to

improve maturation protocols [

2 , 21 ]. In addition to genes encod-

ing proteins related to embryo tolerance to desiccation (LEA,

HSP) ( see Subheading

2.4.6 ), various genes involved in defense

and maintenance of redox homeostasis (oxidative stress) were dif-

ferentially expressed [

2 , 19 , 23 , 25 , 42 , 92 ]. Defense genes were

activated in suboptimal maturation conditions promoted by abi-

otic stresses, such as anoxia. Both alcohol dehydrogenase and pyru-

vate decarboxylase genes were upregulated in P. pinaster EMs

during maturation in unfavorable conditions [

25 ]. Regulation of

2.4.8 Defense Genes

and Maintenance of Redox

Homeostasis

Jean-François Trontin et al.

189

these genes was in agreement with activation of the glycolytic path-

way, but they might also be involved in anoxia tolerance through

alcoholic fermentation as suggested by increased expression of

SuSy3 gene. SuSy genes are responsive to low oxygen level and

promote adequate sugar supply under anaerobic conditions. A

bifunctional enolase 2 gene related to stress-regulated transcrip-

tional networks was concomitantly upregulated, as also observed

during early embryogenesis in unproductive P. abies lines [

23 ]. In

contrast, enolase genes were upregulated in productive lines of P.

radiata [

1 ]. Enolase gene expression may reveal suboptimal matu-

ration conditions (oxygen or other abiotic stress) or, alternatively,

enhanced carbon metabolism as enolase is also involved in glycoly-

sis/gluconeogenesis. Additional regulated genes with a likely

defense function to overcome culture constraints include a puta-

tive cytochrome P450 gene and genes related to biosynthesis of sec-

ondary metabolites. Cytochrome P450 was expressed during early

embryo development in P. radiata [

89 ] and downregulated in P.

abies cotyledonary SE [

23 ]. This gene encodes monooxigenases

involved in plant responses to PGRs (e.g., ABA , IAA) and environ-

mental stresses such as osmotic stress. Pathways related to second-

ary metabolism such as phenylpropanoids or ﬂ avonoids were

overrepresented in the transcriptome of L. kaempferi proliferating

EMs [

24 ]. Flavanone 3-hydroxylase gene ( F3H ) involved in ﬂ avo-

noid biosynthesis was downregulated in P. abies during EM prolif-

eration [

23 ], whereas F3H and several genes related to ﬂ avonol

metabolism were strongly upregulated at the onset of SE matura-

tion in P. pinaster [

25 ]. Activation of ﬂ avanone hydroxylation and

the subsequent production of condensed tannins have been associ-

ated with stress resistance in plants. Surprisingly, genes involved in

response to pathogens are also regulated during embryo develop-

ment in P. abies , particularly during transition from early to late

embryogenesis [

2 ]. A positive regulator ( SN1I ) of the systemic

acquired resistance (SAR) response was expressed during early

somatic embryo genesis in P. radiata [

1 ]. This gene contributes to

the regulation of pathogenesis-related (PR) protein genes and sali-

cylic acid-mediated transduction of the SAR signal.

The maintenance of an efﬁ cient cellular homeostasis by redox

antioxidant metabolites, such as glutathione, is critical to regulate

oxidative stress and the associated production of reactive oxygen

species ( ROS ), free radicals and hydrogen peroxides resulting from

aerobic metabolism [

2 , 19 , 23 , 42 , 92 ]. Redox homeostasis may

represent a generic sensor for controlling embryo development

[

23 ] through PCD activation by ROS [ 49 ] or interplay of glutathi-

one with NADP-linked thioredoxin in the frame of auxin transport

and signaling. Both SE yield and quality are affected by deregula-

tion of glutathione metabolism [

124 ]. Expression of cytosolic ascor-

bate peroxidase and thioredoxin H genes, involved in both

detoxiﬁ cation and control of the cellular redox state, were down-

regulated during EM proliferation in P. abies productive lines [

23 ].

Molecular Aspects of Conifer Embryo Development

190

It was suggested that high oxidative stress could alter the activity

of ascorbate peroxidase in unproductive lines and preclude embryo

development [

125 ]. Metabolic activity in developing P. pinaster

ZE was reﬂ ected by a general overrepresentation of oxidation–

reduction processes with a prevalence of glutathione metabolism

(glutathione thiolesterase activity, expression of glutathione trans-

ferases ), especially during early development [

19 ]. Related genes

were regulated in P. glauca [

92 ] during early SE development

( glutathione-S-transferase , glutathione peroxidase ) at the transition

from pre- to early-cotyledonary embryo ( glutathione reductase )

and at the cotyledonary stage ( glutathione peroxidase , ascorbate

peroxidase ). Genes encoding another well-known antioxidant

enzyme ( superoxide dismutase , SOD ) were similarly upregulated at

the cotyledonary stage. In L. kaempferi expression of a Cu/Zn

SOD gene (plastocyanin) putatively regulated by miR398 increased

at the pre-cotyledonary stage [

24 ]. SOD genes were not differen-

tially expressed at the onset of late SE development in P. pinaster ,

but resulted in enhanced SOD protein production ( see Subheading

3 ). Differential expression of SOD proteins may relate with the

concomitant upregulation of GLP genes. Germins and GLPs are

usually located in the extracellular matrix where they can have both

enzymatic (oxalate oxidase, SOD) and nonenzymatic activities

(auxin-binding protein, serine protease inhibitor), associated with

either response to stress or developmental regulation ([

51 ], and

references therein). Germin and GLP enzymatic activities result in

the production of hydrogen peroxide that may be involved in cell

wall remodeling during stress response s and/or development. Two

GLP genes most similar to a germin with putative oxalate oxidase

activity were speciﬁ cally expressed in proliferating EMs of P. radi-

ata [

89 ]. In L. x marschlinsii , expression of a GLP gene in prolif-

erating EMs ( LmGER1 ) was associated with SOD activity in

apoplastic proteins extracted from early SE [

51 ]. LmGER1 expres-

sion was located in suspensor cells and at the junction with EM

during proliferation and persisted at the embryonal root cap after

transfer to maturation medium. Interestingly, LmGER1 expression

corresponded to the pattern of active PCD during embryo devel-

opment in conifers. Downregulation of LmGER1 in proliferating

EM resulted in reduced SE yield, asynchronous development, and

precluded plantlet regeneration.

3 Proteomics of Conifer Somatic and Zygotic Embryo Development

The development of high-resolution, two-dimensional polyacryl-

amide gel electrophoresis (2D-PAGE), coupled with chromato-

graphic separation and identiﬁ cation through mass spectrometry

(MS), has allowed increased and untargeted qualitative proteome

coverage, together with quantitative measurements of proteins

Jean-François Trontin et al.

191

involved in plant development [ 126 , 127 ]. Various proteome

variations can be expressed by the same or different genomes

according to ontogenetic programs or as a major component of

phenotype plasticity . Many fundamental activities performed by

proteins (especially enzymes) are involved in most metabolic and

signaling pathways. Proteomics therefore aims to identify and

assign physiological functions to “candidate proteins,” contribut-

ing to developmental processes and valuable traits.

Despite signiﬁ cant inputs of quantitative proteomics to molec-

ular identiﬁ cation and functional characterization of embr yogenesis-

related genes in model and crop plants [

20 , 36 ], there are relatively

few recent contributions in conifers (Table

2 ) focused on somatic

[

4 , 25 , 93 , 96 , 100 – 103 ] and/or zygotic embryo genesis [ 95 , 97 –

99 , 104 , 105 ]. First-generation approaches are efﬁ cient, but have

high experimental and technical requirements [

126 , 127 ] and can

be biased towards hydrophilic proteins [

103 ]. New methods such

as 2D difference gel electrophoresis (2D-DIGE) and unbiased 1D

SDS-PAGE combined with isobaric tags for relative and absolute

quantitation (iTRAQ) will considerably facilitate the identiﬁ cation

of differentially expressed proteins and will offer a more global

view of the proteome dynamics [

103 , 104 ]. Moreover, the identi-

ﬁ cation rate of multiple proteins in large proteomic datasets is still

challenging because it largely depends upon the availability of

exhaustive genome resources [

4 , 7 , 101 , 127 ]. This information is

expected to substantially expand according to completed, publicly

available genome sequences [

11 – 13 ].

Published proteomics studies of embryo development in coni-

fers are currently restricted to nine species from the Araucaria ,

Cunninghamia , Cupressus , Larix , Picea , and Pinus genera (Table

2 ) and are mainly focused on late SE or ZE development.

Proteomics considerably enhanced the sensitivity and scale (up to

1000 spots detected per gel) of protein expression studies during

embryo development [

4 , 100 , 102 ]. Previous qualitative or semi-

quantitative methods were often restricted to a few major proteins,

especially storage proteins [

123 , 128 ]. Pioneering investigations of

temporal protein changes in P. abies [

129 ] and Cupressus sempervi-

rens [

100 ] revealed the large sets of protein expression patterns

that can be associated with embryo developmental stages. The

paradigm shift in technology resulting from proteomics was fur-

ther illustrated in P. glauca [

4 ]. Most differentially expressed pro-

teins (79 %) identiﬁ ed in this work were indeed new proteins not

previously associated with embryo development. Biological and

functional relevance of new candidate proteins may be elucidated

and ultimately provide opportunities for reﬁ ning the somatic

embryo genesis process.

Here, we brieﬂ y review the importance of speciﬁ c protein

functional classes that were either validated ( storage proteins ) or

reinforced (metabolic/cellular processes, stress-response proteins)

by proteomics studies of embryo development .

Molecular Aspects of Conifer Embryo Development

192

Developing SEs and ZEs in conifers have been shown to accumu-

late major storage proteins of the globulin (legumin, vicilin) and

albumin families based on electrophoretic mobility patterns. These

assumptions were validated by MS approaches in P. strobus [

128 ],

P. pinaster [

105 ], P. glauca [ 4 ], P. abies [ 93 ], and L. x eurolepis

[

102 ]. Expression pattern of storage proteins was similar in SE and

ZE [

4 , 102 , 105 , 128 ], reaching a maximum at the cotyledonary

stage. Storage proteins were already detected at the late stage of

early embryogenesis ( P. glauca ) or at the pre-cotyledonary stage

( L. x eurolepis ). Protein accumulation and SE growth are affected

by maturation duration and cultural conditions [

93 , 128 ]. Routine

tracking of the main storage proteins may be valuable for assessing

the quality of matured embryos. The most dominant vicilin-like

storage proteins have been proposed as markers of SE develop-

ment in P. glauca [

4 ] and L. x eurolepis [ 102 ]. Similarly, three

vicilin- and legumin-like proteins as well as two cupin domain-

containing storage proteins were identiﬁ ed in P. pinaster as candi-

date biomarkers for the late cotyledonary SE/ZE stage [

105 ].

The activation of various metabolic and cellular processes during

SE development could be emphasized by proteomic data. A com-

parison of immature and mature embryos in L. x eurolepis showed

an increase in proteins involved in primary metabolism (glucose,

pentose, starch), suggesting active glycolysis, nucleotide metabo-

lism, and accumulation of storage carbohydrates [

102 ]. However,

in both L. x eurolepis [

101 ] and P. pinaster [ 25 ], the glycolytic

pathway appeared to be reduced under favorable maturation con-

ditions (high gellan gum concentration). Reduced water availabil-

ity induced by high gellan gum may promote a decrease in carbon

catabolism through downregulation of key proteins involved in

glucose or pentose metabolisms. The decreased level of glycolysis

in EM cultivated on favorable maturation medium has been associ-

ated with increased embryo dry weight ( L. x eurolepis , P. pinaster )

and enhanced starch accumulation ( P. pinaster ) at both cytological

and proteomic levels (e.g., upregulation of glucose-1-phosphate

adenylyltransferase). Upregulation of proteins involved in amino-

acid metabolism was also highlighted in Larix [

102 ] at the mature

embryo stage, and is indicative of active protein synthesis as

observed during zygotic embryo genesis in other conifers [

95 , 98 ,

99 , 104 ]. Accordingly, the maturation treatment was reported to

induce changes in nitrogen metabolism in mature embryos of P.

abies [

93 ] through differential expression of key enzymes for

glutamine , glutamate and arginine synthesis. In accordance with

gene expression studies ( see Subheading

2.4.7 ), differential expres-

sion of various proteasome subunits in P. pinaster [

25 ] and P. abies

[

4 ] as well as elongation factor II protein during EM proliferation

in A. angustifolia [

96 ] supported the importance of controlled

proteolysis and protein synthesis when embryos are stimulated to

3.1 Storage Proteins

3.2 Proteins Involved

in Metabolism

and Cellular Processes

Jean-François Trontin et al.

193

develop. Lippert et al. [ 4 ] proposed the proteasome complex as a

source of protein markers to evaluate embryo development .

Teyssier et al. [

102 ] also suggested that various differentially

expressed proteins from the primary and amino-acid metabolisms

are suitable targets for marker validation.

Other metabolic pathways with important roles in embryogen-

esis were also suggested to be activated as a result of enhanced

amino-acid metabolism, especially methionine and SAMet synthe-

sis. A SAMet synthetase was found to be upregulated in L. x euro-

lepis mature SE [

102 ], expressed at various developmental stages

of P. glauca SE [

4 ] and expressed from proembryogeny to late ZE

stages in A. angustifolia [

95 ]. This protein is involved in DNA

methylation, polyamines and ethylene biosynthesis. Proteome

analysis in A. angustifolia revealed a set of ten proteins unique to

eight responsive or two recalcitrant lines to maturation treatment

[

96 ]. It is suggested that embryogenic potential could be associ-

ated with upregulation of SAMet synthetase during EM prolifera-

tion. Interestingly, Jo et al. [

96 ] provided data showing increased

ethylene release and lower putrescine content in responsive lines.

Other important cellular processes upregulated during normal

embryo development included cell wall deposition and cell expan-

sion in P. pinaster (e.g., expansin S2/B14) [

25 ], L. principis-

rupprechtii (e.g., α-1,4-glucan protein synthase) [

103 ], and P. abies

(e.g., reverse glycosylating protein RGP-1) [

4 ], nucleocytoplasmic

transport (e.g., tubulin beta-2 chain, GTP-binding nuclear proteins

Ran-A1) in P. pinaster [

25 ], regulation of membrane trafﬁ cking

(e.g., ADP-ribosylation factor GTPase-activating proteins) in L.

principis-rupprechtii [

103 ], or energy metabolism in A. angustifolia

(e.g., mitochondrial ATPase beta subunit) [

96 , 97 ] and P. abies

(e.g., ATP synthase, H+ transportin) [

4 ]. ATP production and

catabolism have been associated to competent embryo maturation

and structural reorganization via PCD . In P. pinaster , active PCD

was revealed by combined analysis of transcriptomic and proteomic

datasets, showing upregulation of both chitinases and disulﬁ de

isomerase [

25 ]. A nondefensive role of chitinase IV in early SE

development was also supported in L. principis-rupprechtii [

103 ].

Available proteomics studies emphasized the omnipresent

“background” expression of stress-related protein s during SE

development and maturation. This is in accordance with the data-

sets provided by transcriptomic and metabolomic studies ( see

Subheadings

2 and 4 , respectively). Such proteins represented up

to 6.7 % of differentially expressed proteins in P. abies [

93 ] and are

mainly involved in response to oxidative stress, anoxia, prevention

of apoptosis, and tolerance to cellular dehydration. Oxidative stress

may be induced by water and/or osmotic stress and it results in

production of ROS , ATP depletion and, ultimately, in apoptosis

[

130 ]. The SOD enzyme involved in detoxiﬁ cation processes

3.3 Stress-Related

Proteins

Molecular Aspects of Conifer Embryo Development

194

through regulation of oxidative stress was found overexpressed in

early developing SE in P. pinaster [

25 ] and in mature SE in L. x

eurolepis [

102 ]. A similar pattern was observed during ZE develop-

ment in P. massoniana [

104 ]. A GLP was also overexpressed in P.

pinaster [

25 ], suggesting active antioxidant protein production.

The interest of GLPs as predictive markers of embryo development

is well supported by proteomics ([

25 ], and references therein).

Accordingly, suboptimal conditions for embryo maturation in L. x

eurolepis resulted in upregulation of SOD and activation of second-

ary metabolism enzymes, possibly to cope with increased produc-

tion of free radicals [

101 ]. Similarly, Jo et al. [ 96 ] revealed that

NADH dehydrogenase in A. angustifolia was upregulated in one

recalcitrant line to a maturation treatment, thus suggesting a dis-

turbed cell redox system. NADH dehydrogenase is a component

of the plant energy-dissipating mitochondrial system preventing

excessive ROS production. ROS were recently revealed as impor-

tant signaling molecules for activation of PCD and normal SE

development in L. leptolepis [

49 ]. Overexpression of catalase (anti-

oxidative enzyme) in non-embryogenic callus compared to EMs

provided indirect evidence in L. principis-rupprechtii for excessive

ROS generation in response to culture conditions [

103 ].

Abiotic stress may also result from anaerobic conditions during

in vitro culture. As previously discussed, enolase is involved in gly-

colysis/gluconeogenesis pathways but can also be induced by abi-

otic stresses such as oxygen levels. Enolase accumulates in P. glauca

mature embryos and has been proposed as a putative protein

marker of normal embryo development [

4 ]. In L. x eurolepis , two

enolase isoforms were found overexpressed after a suboptimal mat-

uration treatment [

101 ]. Overexpression in P. glauca of a

submergence- induced protein at the early SE stages was inter-

preted as a possible response to oxygen stress promoting cell elon-

gation in developing embryos. Several enzymes involved in the

glycolytic pathway (e.g., alcohol dehydrogenase, pyruvate decar-

boxylase) were similarly activated under unfavorable SE matura-

tion conditions in P. pinaster [

25 ]. Interestingly, both alcohol

dehydrogenase and pyruvate decarboxylase expressions were

recently reported to be involved in tolerance to anoxia [

131 ].

Various protein families with important protective roles during

abiotic stresses resulting in cellular dehydration were conﬁ rmed to

accumulate in mature embryos, including LEAs and group 2 LEAs

(dehydrins), HSPs and small HSPs. Members of these protein fam-

ilies were upregulated in P. abies embryos matured with sucrose as

a carbon source [

93 ]. The presence of 3 % sucrose signiﬁ cantly

improved SE germination rate by promoting the acquisition of

desiccation tolerance. LEA and dehydrins were reportedly shown

to accumulate in plants during late embryogenesis. In L. x eurolepis

[

102 ], a set of 21 proteins annotated as belonging to HSPs or

related to protein folding were found differentially expressed in

Jean-François Trontin et al.

195

developing versus mature embryos. Most were upregulated at the

mature stage in accordance with the proposed role of HSPs in cel-

lular protection (protein stabilization and refolding). HSPs and

small HSPs were similarly detected in mature SE of P. abies [

93 ]

and were overexpressed in both cotyledonary SE and maturing ZE

in P. pinaster , together with various LEAs [

105 ]. HSPs were also

found to accumulate at early SE stage in L. principis-rupprechtii

[

103 ], from early to mature SE stages in L. x eurolepis [ 102 ] and P.

glauca [

4 ], as well as during ZE development in Cunninghamia

lanceolata [

99 ]. HSP expression is known to be induced by ABA

and there is strong evidence that these proteins are required

throughout embryogenesis from initiation to early seedling growth

([

102 ], and references therein). HSPs and other stress-related pro-

tein s are also overexpressed during maturation in suboptimal con-

ditions [

101 ]. Proteomics therefore strengthened both the

protective function of HSPs in response to abiotic stress and their

ubiquitous role in protein folding, assembly translocation and deg-

radation during embryo development .

4 Metabolomics of Conifer Somatic and Zygotic Embryo Development

Metabolite proﬁ ling can be achieved in plants with high resolution

and good sensitivity by using gas chromatography coupled with

mass spectrometry (GC/MS) or nuclear magnetic resonance

(NMR) spectroscopy. Both GC/MS and NMR spectroscopy are

high-throughput techniques for unbiased acquisition of quantita-

tive and qualitative data on multiple metabolites. These technolo-

gies are also suitable for time-series studies. Among all “omics,”

metabolomics is considered to provide the most functional infor-

mation since metabolites are the end products of the cellular

machinery. Multivariate data analyses are required to determine

whether combined abundances of a set of metabolites can be asso-

ciated with a speciﬁ c physiological state. Metabolomics is an effec-

tive and increasingly popular approach in conifers for monitoring

physiological responses to environmental variation [

132 ].

Applications to embryo development are currently scarce (Table

2 )

because of the technical requirements involved and the integrated

proteomics information (identiﬁ cation of enzyme substrates)

needed to fully interpret the data [

133 ].

Interestingly, metabolic proﬁ ling already provided relevant

information in Picea species about the biochemical status of EMs

at, or during, transition between different embryo development al

stages and in response to different maturation conditions [

5 , 93 ,

94 ]. The metabolic signature has also been demonstrated in P.

taeda to accurately predict the ability of proliferating EMs to

regenerate SE [

6 ]. Therefore, it is expected that these studies will

provide not only a better understanding of SE development, but

Molecular Aspects of Conifer Embryo Development

196

also tools for monitoring early metabolic events determining SE

physiology. Metabolite proﬁ ling can be used to analyze intracellu-

lar metabolites ( metabolic ﬁ ngerprinting ) or, alternatively, the

metabolite composition of fresh and spent culture medium ( meta-

bolic footprinting ). The latter noninvasive method is not affected

by rapid turnover of intracellular metabolites and is likely to yield

valuable information about critical metabolites, especially for the

complementation and interpretation of metabolic ﬁ ngerprints [

6 ,

37 ]. Metabolic footprinting was performed in P. glauca (NMR

spectroscopy) to identify signiﬁ cant metabolites (35 compounds

detected) involved in SE proliferation and maturation [

5 ]. Strong

evidence for divergent metabolic processes and different EM phys-

iological state in proliferation and maturation media was obtained

within 48–72 h. Major sources of metabolic variation in culture

media over time included carbohydrates , amino acids (consump-

tion of medium compounds), and also processed metabolites

excreted by the cultured cells. Early sucrose hydrolysis and prefer-

ential use of glucose over fructose by embryogenic cells was appar-

ent in both conditions. Most other discriminating metabolites

were overrepresented in the proliferation medium and were indica-

tive of storage protein synthesis and regulation, nitrogen transport

and ammonium assimilation (5-oxoproline, glutamine , BCAA/

branched chain amino acids), response to various stresses and

intracellular/inter-organ signaling (GABA/γ-aminobutyric acid),

biosynthesis of phenylpropanoid compounds (phenylalanine) and

cell expansion (malate). BCAA and GABA proﬁ les are particularly

suggestive of a metabolic imbalance as a result of altered coenzyme

A biosynthesis during maturation. Such a metabolomic-generated

hypothesis paves the way for expression studies of speciﬁ c genes

involved in this pathway in conifers.

In P. abies , metabolic ﬁ ngerprinting (GC/MS) was used to

study metabolic events involved in normal SE development [

94 ].

Three different embryogenic lines with blocked, aberrant, or nor-

mal phenotype were investigated. Signiﬁ cant metabolites were

identiﬁ ed from EM in proliferation through to cotyledonary

embryos. Sucrose was revealed as the main carbohydrate in prolif-

erating EM, whereas maltose was signiﬁ cant during late embryo-

genesis in the normal line. In contrast, a preponderance of fructose

was observed in lines with abnormal phenotypes. Metabolite pro-

ﬁ ling therefore conﬁ rmed previous data showing that supplemen-

tation of maturation medium with maltose to promote nutritional

stress (cellular carbon restriction) could improve embryo develop-

ment ([

134 ], and references therein). This hypothesis could be

partially veriﬁ ed in P. abies as maturation yield increased when

sucrose was replaced by a combination of maltose and PEG [

93 ].

However, the latter formulation was detrimental to SE germina-

tion . Metabolic ﬁ ngerprinting could separate samples according to

maturation condition (45 compounds detected) and revealed that

Jean-François Trontin et al.

197

SE treated with maltose and PEG accumulated less rafﬁ nose. The

metabolite signature therefore suggested that poor germination

rate results from reduced content in rafﬁ nose family oligosaccha-

rides (RFOs) that are involved in the acquisition of desiccation

tolerance together with sucrose and LEA.

Evidence suggesting that metabolic response to osmotic stress

may be a key factor involved in normal embryo development was

gained through metabolic ﬁ ngerprinting of P. taeda proliferating

EMs [

6 ]. In this large study that detected 208 metabolites,

embryogenic culture’s regenerative capacity was not only inﬂ u-

enced by the genetic background and maturation conditions, but

also by the metabolic status of the proliferating culture at the time

of sampling. It appeared that a culture containing developmentally

advanced immature embryos is more likely to produce cotyledon-

ary SE, as previously observed in other pine species [

134 ]. Among

the 47 identiﬁ able metabolites selected to build a descriptive model

of cell line ability to regenerate SE, several were related to osmo-

protectants. Proline, serine and arabitol contents may be indicative

of biological stress during proliferation as a negative relationship

with culture productivity was observed. In contrast, a positive cor-

relation was found with sorbitol accumulation, suggesting that

some osmoprotective compounds may also play a role in prevent-

ing biological stress and preserving culture responsiveness to mat-

uration treatments. The method was therefore highly efﬁ cient at

identifying both informative metabolites and their relationships to

gain insights into the transition from immature to mature SE. It is

based on the multivariate analysis of a metabolite subset selected

through a stepwise modeling procedure following the Bayesian

information criterion. In addition, the model was demonstrated to

accurately predict the regenerative capacity of proliferating EM in

a genotype-independent manner. A robust assay based on multiple

predictor metabolites accounting for genetic variability could

prove invaluable in pine as the regenerative capacity is invariably,

although erratically, decreasing as a function of line aging [

15 ].

5 Conclusion and Future Directions

The molecular biology of conifer embryo development has begun

to beneﬁ t from genome-wide approaches. Technical requirements

are still high [

22 , 132 , 135 ] and there are also strong limitations to

the interpretation of these large datasets. The development of

comprehensive genome resources [

10 – 13 ] is expected to consider-

ably increase the identiﬁ cation rate of differentially expressed

genes. The “holy grail” will then be to characterize the function

and molecular regulation of important genes in metabolic net-

works to model embryo development through integration of tran-

scriptomic, proteomic and metabolomic data [

20 , 37 , 38 , 133 ].

Molecular Aspects of Conifer Embryo Development

198

Such a systems biology approach is likely to provide tested clues for

the development of somatic embryo genesis in plants, including

conifers.

“Omics” has started to improve our knowledge of conifer

embryo development . Transcriptome proﬁ ling of embryogenesis-

related genes in conifers has shown high homology with model

angiosperms . It is suggested that differences in the molecular regu-

lation of embryogenesis may mainly arise from spatiotemporal

variation in gene expression. Several important processes are appar-

ently conserved in plants [

2 , 19 ], in particular early organization of

apical-basal embryo patterning driven by polar auxin transport and

activation of the auxin-mediated response machinery during late

embryogenesis (radial embryo patterning). Conserved expression

proﬁ les were also revealed for important epigenetic regulator s

( chromatin remodeling ) involved in temporal and organ-speciﬁ c

expression of homeotic genes [

19 ]. Transcriptomic studies have

highlighted the complexity of processes and genes involved in the

spatiotemporal development of embryos, from embryogenesis

induction [

20 , 21 ] to the switch from embryonic to vegetative

growth [

2 , 23 , 25 , 92 ]. A reference gene regulation network has

been proposed for embryogenesis induction in plants [

20 ], but

there is a need for dedicated studies in conifers to further elucidate

these pathways [

21 , 70 , 76 ]. Transcriptome proﬁ ling and, to a

lesser extent, proteomics have revealed multiple genes associated

with early embryo and late embryo development. An impressive

picture of coordinated functions and genes has been obtained dur-

ing development of SE in P. abies [

2 , 23 , 92 ], and of ZE in P.

pinaster [

19 ]. TFs genes appeared to have central roles in spatio-

temporal modulation of both auxin- and GA-mediated responses,

especially during early embryogenesis (e.g., LEC and AGAMOUS ).

Later during development, LEC gene s and other master regulators

revealed in A. thaliana ( ABI3 and FUS3 ) are likely to have similar

roles in conifers, i.e., induction of ABA -dependent response that

may modify EM responsiveness to auxin and GA, but also to other

signaling molecules ( polyamines , ethylene ). Regulation of these

pathways could be involved in the developmental switch from

embryonic to vegetative growth. Various additional processes have

been suggested to have general functions in development stages

such as PCD , megagametophyte signaling, cell wall modiﬁ cation,

epigenetic regulation (DNA methylation, small RNAs), carbohy-

drate, protein or energy metabolisms, and response to stress.

Opportunity for modulation of any of these pathways could be of

practical interest to reﬁ ne speciﬁ c steps of seed production or

somatic embryo genesis in conifers.

Genome-wide proﬁ ling offers the possibility to check the qual-

ity of proliferating EMs and developing SE at the molecular level

with unprecedented accuracy and throughput, showing that omics

is already providing some important clues to improve conifer

Jean-François Trontin et al.

199

embryo development . Early molecular screening can help prevent-

ing unnecessary expenses associated with EMs cultivated in unfa-

vorable conditions and/or with low ability to be converted into

high quality plantlets. There is a choice of proposed marker genes

revealing speciﬁ c processes and adaptation at each developmental

stage or transition, from embryogenesis induction and initial dem-

onstration of embryogenecity [

21 , 26 , 70 , 76 , 89 ], to early embryo

patterning [

1 , 2 , 58 , 65 , 90 ] and late embryo development [ 19 ,

23 , 25 , 52 , 77 , 78 , 84 , 89 ]. Substantial support has been obtained

for a few proteins proposed as robust markers of embryo develop-

ment in P. glauca (vicilin, enolase, proteasome subunit) [

4 ], L. x

eurolepis (vicilin) [

102 ] and P. pinaster (GLP, ubiquitin-protein

ligase) [

105 ]. More pragmatically, a selected subset of metabolites

has been demonstrated in P. taeda [

6 ] to accurately predict in a

genotype-independent way the ability of proliferating embryo-

genic lines to regenerate cotyledonary SE.

There is also a large set of genes involved in epigenetic regula-

tion that were repeatedly highlighted in transcriptomic studies

with high relevance for proper embryo development [

1 , 19 , 23 ,

50 , 92 ]. In particular, expression of various miRNAs with stage-

speciﬁ c modulation was associated with the regulation of impor-

tant genes during somatic embryo genesis in L. kaempferi [

50 ],

including genes involved in the regulation of auxin-mediated

response, cell wall modiﬁ cation, embryo pattern formation, ABA

response, oxidative stress and miRNA biogenesis. Some miRNA

appeared to have functions in maintaining the embryogenic

potential ( miR159 , miR171 ). It would therefore be interesting to

infer the general signiﬁ cance of embryogenesis-related miRNAs in

conifers.

Although SEs develop without true megagametophyte [

2 ],

there is a consensual trend toward approaching “substantial equiv-

alence” of SEs with ZEs [

26 , 42 , 77 , 78 , 91 , 92 ]. Comparative

“omics” of SE and ZE is a promising tool to elaborate new

strategies to reach the performance standard of seedlings.

Transcriptomic proﬁ ling in P. taeda [

27 ] and proteomic analysis

in P. pinaster [

105 ] gave strong evidence that gene expression in

cotyledonary SE obtained after “appropriate” culture time in

“reﬁ ned” maturation conditions did not conform to that of fully

mature ZE, but to that of earlier, immature cotyledonary stages.

Similar conclusions were made in P. pinaster after a study of genes

involved in nitrogen metabolism and chloroplast development

[

78 ]. Data analysis suggested speciﬁ c protocol reﬁ nements at

either the maturation or post-maturation step. Optimization of

SE maturation in P. taeda resulted in similar expression patterns of

genes involved in controlled proteolysis and synthesis of storage

proteins compared with ZE [

77 ]. In A. angustifolia , comparative

transcriptomics of SE and ZE revealed auxin signaling failure dur-

ing SE development [

91 ].

Molecular Aspects of Conifer Embryo Development

200

Another way to estimate if embryo quality could be enhanced

is to compare different maturation conditions. In P. glauca , the

beneﬁ cial effect of PEG in the maturation medium (improved SE

yield and quality) could be demonstrated using a cDNA array strat-

egy [

92 ]. This study provided the ﬁ rst evidence that transcriptome

proﬁ ling could predict embryo quality, as many regulated genes

between PEG-treated and control lines were identiﬁ ed in early

developing embryos. A similar approach integrating both tran-

scriptomic and proteomic proﬁ ling was implemented in P. pinaster

to study the early molecular events involved in SE development

promoted by high gellan gum [

25 ]. Differential expression of

genes associated with embryo development or culture adaptive

responses, as early as 1 week after exogenous ABA treatments, sup-

ported integrated genome-wide proﬁ ling as a robust diagnostic

and predictive tool for detecting disruption of critical pathways for

normal SE development. Interestingly, gene expression studies in

P. abies have already inﬂ uenced protocol improvement (accelerated

and synchronized SE development) through either modiﬁ cation of

maturation conditions (latrunculin B treatment affecting actin

gene expression) [

54 ] or genetic engineering of proliferating EM

( HBK3 overexpression promoting AGO upregulation) [

55 ].

It is foreseeable that genome-wide proﬁ ling will be further

implemented in both important species (to achieve cost-effective SE

variety deployment) and orphan species (to save labor and associated

cost of development). Integrating transcriptomic and proteomic

approaches may inherently offer robust tools to assess embryo devel-

opment [

25 ]. Metabolomics may also provide unique opportunities

for constructing genotype-independent, predictive models of

embryogenesis-related traits. Interpretation of “omic” data may

help identify new directions for gene expression proﬁ ling of selected

metabolic pathways underpinning embryo development.

Acknowledgements

The preparation of this chapter was supported through various

projects funded by (1) the French National Research Agency

(GENOQB: ANR-05-GPLA-027, SUSTAINPINE: ANR-09-

KBBE-007, XYLOFOREST: ANR-10- EQPX-16), (2) the

European Community’s Seventh Framework Programme

(FP7/2007-2013, Grant Agreement n° 289841-PROCOGEN),

and (3) the French Regional Councils of “Région Centre”

(EMBRYOME: 33639, IMTEMPERIES: 2014- 00094511) and

“Région Aquitaine” (EMBRYO2011: 09012579- 045). K.K. was

supported by Natural Resources Canada, Canadian Forest Service.

Mrs. Isabelle Lamarre (NRCan, CFS) is thanked for English

editing.

Jean-François Trontin et al.

201

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_9, © Springer Science+Business Media New York 2016

Chapter 9

Androgenesis in Solanaceae

Jose M. Seguí-Simarro

Abstract

The Solanaceae is one of the most important families for global agriculture. Among the different solana-

ceous species, tobacco ( Nicotiana tabacum ), potato ( Solanum tuberosum ), tomato ( Solanum lycopersi-

cum ), eggplant ( Solanum melongena ), and pepper ( Capsicum annuum ) are ﬁ ve crops of outstanding

importance worldwide. In these crops, maximum yields are produced by hybrid plants created by crossing

pure (homozygous) lines with the desired traits. Pure lines may be produced by conventional breeding

methods, which is time consuming and costly. Alternatively, it is possible to accelerate the production of

pure lines by creating doubled haploid (DH) plants derived from (haploid) male gametophytes or their

precursors (androgenesis). In this way, the different steps for the production of pure lines can be reduced

to only one generation, which implies important time and cost savings. This and other advantages make

androgenic DHs the choice in a number of important crops where any of the different experimental

in vitro techniques (anther culture or isolated microspore culture) is well set up. The Solanaceae family is

an excellent example of heterogeneity in terms of response to these techniques, including highly respond-

ing species such as tobacco, considered a model system, and tomato, one of the most recalcitrant species,

where no reliable and reproducible methods are yet available. Interestingly, the ﬁ rst evidence of androgen-

esis, particularly through in vitro anther culture, was demonstrated in a solanaceous species, Datura

innoxia . In this chapter, we report the state of the art of the research about androgenic DHs in Solanaceae,

paying special attention to datura, tobacco, potato, tomato, eggplant, and pepper.

Key words Anther culture , Datura , Doubled haploid s , Eggplant , Microspore culture , Microspore

embryogenesis , Pepper , Potato , Tobacco , Tomato

1 Introduction

It is well known that crop productivity can be increased through

the use of hybrids, made by crossing homozygous (pure) lines with

deﬁ ned traits. These lines are traditionally generated by techniques

based on classical breeding, through successive rounds of selﬁ ng

and selection. This requires a considerable amount of time and

resources. However, in recent years alternative techniques, by far

more advantageous than traditional methods, are being used in

some species. These techniques, based on androgenesis , produce

pure, doubled haploid (DH) lines through in vitro regeneration of

plants from microspore / pollen -derived embryos or callus.

210

This experimental pathway, alternative to normal pollen devel-

opment, was discovered 45 years ago by Guha and Maheshwari

[

1 ]. In this route, the pollen grain precursors deviate from the

gametophytic pathway and are in vitro induced to form haploid

embryos or calli [

2 ]. Then, plants can be directly regenerated by

microspore -derived, haploid embryogenesis, or indirectly from an

intermediate haploid callus phase. These plants will be DH if they

duplicate their original haploid genome, or just haploid as the orig-

inal microspore. In the latter case, additional treatments to pro-

mote genome doubling are needed [

3 ]. In both cases, the resulting

plants will have a genetic background exclusively coming from

donor (male) plant, and 100 % homozygous . In other words, they

will be pure lines. From the standpoint of plant breeding, this

alternative reduces the typical 7–9 inbreeding generations neces-

sary to stabilize a hybrid genotype to only one. It is therefore much

faster and cheaper, and obviously this is the main advantage of DH

technology in the context of plant breeding. Within this same con-

text of plant breeding, DHs (homozygous for all of their loci) con-

stitute a valuable tool for the study of the genetic basis of

quantitative traits, including the genetic mapping of complex char-

acters such as production or quality, the most agronomically inter-

esting, and difﬁ cult to be addressed by other approaches. In fact,

they have been successfully used in several crops for breeding plants

with useful agronomic traits such as high yield, earliness, abiotic

stress tolerance, and disease resistance, among others. DHs are also

a powerful tool in transgenesis, to avoid hemizygotes and save time

and resources in the production of plants transformed with the

transgene in both homologous chromosomes. Moreover, from a

scientiﬁ c point of view, these lines are also very useful for basic

studies of linkage and estimation of recombination fractions. They

are also an extremely useful tool for genetic selection and screening

of recessive mutants, because the phenotype of the resulting plants

is not affected by the effects of dominance, and recessive pheno-

types can be unmasked. For example, recessive embryo-lethal

genes would be expressed in haploid embryos, and thereby elimi-

nated for future generations.

Microspore -derived embryos (MDEs) or calli can be obtained

using two main technical approaches based on in vitro culture:

anther culture and isolated microspore culture . Anther culture is

the easiest option. It consists on the excision of anthers from the

ﬂ ower bud, followed by their in vitro culture in a generally semi-

solid, agar -based culture medium. After few weeks, microspores in

the pollen sac transform into MDEs or calli and emerge out of the

anther, which becomes necrotic. This approach is relatively fast and

inexpensive, compared to the other option. This is why anther

culture is the most adopted method to produce androgenic DHs.

However, this method does not exclude the occasional appearance

of somatic embryo s from anther tissues, or the uncontrollable

secretory effect of the tapetal layer surrounding the pollen sac,

Jose M. Seguí-Simarro

211

which prevents us from having a strict control of the culture condi-

tions. In addition, anther cultures need weeks-months to produce

MDEs, and have a limited efﬁ ciency, generally producing only a

few embryos per cultivated anther. All these limitations can be

overcome by the direct isolation and culture of microspores, which

is a more technically demanding method, but it is faster and more

efﬁ cient. In some herbaceous species, where isolated microspore

cultures are well set up, in 1–3 weeks it is possible to get hundreds

or even thousands of embryos from the microspores contained in

a single anther. It is evident that, where possible, microspore cul-

tures are largely preferred over anther cultures. Using either anther

or microspore cultures, at present there are systems for DH pro-

duction in few hundreds of species of agronomic interest, from

herbaceous crops such as wheat , barley , rice, rapeseed ( canola ),

tobacco , or corn to trees such as clementine , mandarin, or cork,

among others (reviewed in refs. [

4 – 7 ]). However, except for model

species such as rapeseed, barley, or tobacco, the efﬁ ciency is still

very low. This is even more critical in horticultural crops of high

agronomic interest, like those belonging to the Solanaceae family.

The Solanaceae family of ﬂ owering plants comprises between

3000 and 4000 species in about 90 genera [

8 ]. Among these gen-

era, the largest is Solanum L., estimated to contain 1500 species

[

9 ], and nearly 50 % of the diversity of the Solanaceae family. This

family is one of the most important in terms of agricultural inter-

est, and includes ﬁ ve major cultivated crop plants [

10 ], namely

potato ( Solanum tuberosum ), pepper ( Capsicum annuum ), egg-

plant ( Solanum melongena ), tobacco ( Nicotiana tabacum ) and

tomato ( Solanum lycopersicum ). This family also includes the wild

relatives of these ﬁ ve species, as well as many other plants belong-

ing to the genus Datura , ornamentals such as those of the genus

Petunia , or toxic, poisonous plants such as mandrake ( Mandragora

ofﬁ cinarum ), henbane ( Hyoscyamus niger ), or deadly nightshade

( Atropa belladonna ). In Solanaceae, ﬂ owers are typically conical or

funnel-shaped, with ﬁ ve petals, usually fused, persistent sepals, and

a general ﬂ oral formula of K(5)[C(5)A5]G(2). Stamens are bithe-

cate and usually longitudinally or poricidally dehiscent. The ovaries

are superior and biloculate. Seeds are usually small, round and ﬂ at.

Fruits are in general berries, as in tomato or wolfberry, or drupes

or dehiscent capsules, as in the genus Datura . Most Solanaceae

have a basic chromosome number of x = 12, being most of them

diploid (2 n = 2 x = 24). Examples of this include tomato, eggplant,

and pepper. However, there are some cases where this number has

increased due to polyploidy. For example, some wild potato

relatives range from diploid to hexaploid (3 n = 6 x = 72), while the

cultivated species of S. tuberosum is allotetraploid (2 n = 4 x = 48),

and the cultivated species of N. tabacum is autotetraploid. The

Solanaceae are a typical example of an ethnobotanical family, mean-

ing that it is extensively exploited and utilized by humans since the

beginning of the agricultural age. It is an important source of food

Androgenesis in Solanaceae

212

and spices, mostly from the agricultural crops, but it is also a source

of medicines and bioactive compounds of pharmaceutical interest,

due to the presence of alkaloids in most plants of this genus. For

example, nicotine in tobacco, or scopolamine, atropine, and hyo-

scyamine in species of the genus Hyoscyamus , Datura , and Atropa .

From a nutritional point of view, the most important crop of this

family is the potato ( S. tuberosum ), where the carbohydrate-rich

tubers are used as food for human and animal nutrition, and as a

source of starch for industrial purposes. In many other Solanaceae,

the fruits are the interesting, edible part of the plant (for example

tomatoes, tomatillos, eggplants, uchuva, sweet and hot peppers).

In view of this, it is not surprising that from an agronomical point

of view, the solanaceous crops are among the most important in

the world.

Despite the tremendous importance of this family for world’s

agriculture, DH technology is not yet efﬁ ciently implemented in

some of these interesting crops. Curiously, the ﬁ rst observation of

in vitro, microspore -derived androgenesis was reported in a solana-

ceous plant by Guha and Maheshwari [

1 ], who described the for-

mation of microspore-derived plants within in vitro-cultured

anthers of Datura innoxia . However, of the ﬁ ve major solanaceous

crops ( pepper , tobacco , potato , eggplant , and tomato ), at present

only in tobacco enough progress has been made to consider this

species as a model system for the study of microspore embryogen-

esis . The rest of interesting solanaceous crops (potato, tomato,

eggplant, and pepper) are still far from the efﬁ ciency achieved in

tobacco or in rapeseed, another model dicot species. In potato,

eggplant, and pepper, only anther culture s seem to work efﬁ ciently

in certain cultivars, but, up to now in tomato, not a single method

has been demonstrated to work efﬁ ciently. Despite of the genetic

proximity of these ﬁ ve species, they seem to respond to induction

very differently. In this chapter we revise the most relevant work

performed in the last four decades pertaining to the study of the

experimental induction of androgenesis through anther cultures

and isolated microspore culture s and the development of the DHs

embryos, calli and plants in datura , the “pioneer” of microspore

embryogenesis, and in the ﬁ ve most important solanaceous crops,

tobacco, potato, tomato, eggplant, and pepper, illustrating when

needed with examples of the practical applications of this technol-

ogy in genetic breeding.

2 Datura

Within the family Solanaceae , the genus Datura comprises several

species (commonly daturas ), widely distributed throughout the

globe, and characterized by their erect or spreading, trumpet-

Jose M. Seguí-Simarro

213

shaped ﬂ owers, and by their spiny capsular fruits that open at

maturity to release numerous seeds. Datura species are herba-

ceous, leafy annuals and short-lived perennials which can reach up

to 2 m in height. Daturas belong to the classic “witches’ weeds”,

along with deadly nightshade, henbane, and mandrake, because

they all contain, primarily in their seeds and ﬂ owers, toxic and hal-

lucinogenic tropane alkaloids such as scopolamine, hyoscyamine,

and atropine. Because of the presence of these substances, Datura

has been used for centuries in some cultures as a poison and as a

key ingredient of love potions and brews. Nowadays, the con-

trolled use of some of these alkaloids at low doses has been adopted

by medicine as treatments for a wide range of diseases. Aside of

their traditional and medical uses, datura plants deserve a honoriﬁ c

position in the ﬁ eld of haploid research for two main reasons. First,

Datura stramonium was the ﬁ rst ﬂ owering plant for which cyto-

logical proof of the discovery of a haploid individual was obtained

[

11 ]. Second, and most importantly, Datura innoxia was the ﬁ rst

plant producing embryos from the microspores contained in their

anthers, when inoculated in a culture dish [

1 ]. Sipra Guha and

Satish C. Maheshwari, two Indian researchers, at that time work-

ing at the University of Delhi (India), ﬁ rst performed such culture

and now they are considered as the founding fathers of haploid and

DH technology [

12 ] ( see Note 1 ).

Since D. innoxia was the ﬁ rst species to produce haploid

MDEs, it is easy to conceive that this experimental system was one

of the ﬁ rst used to investigate such an emerging experimental pro-

cess. Thus, some researchers focused on improving culture condi-

tions in order to increase the “embryogenic power” of pollen in

Datura innoxia [

13 – 15 ]. Other groups focused on the study of the

changes undergone by the microspore within the anther as a con-

sequence of the induction [

16 – 20 ]. Later on, an established proto-

col for microspore embryogenesis in this species permitted the

combination of this technique with others, such as Agrobacterium -

mediated genetic transformation [

21 ], or plant regeneration

through embryogenesis from cultured cells coming from andro-

genic calli [

22 ]. Aside of D. innoxia , other members of the Datura

genus have been used to successfully induce microspore embryo-

genesis, or to study the changes associated to the induction. These

species include D. ferox [

23 ], D. metel [ 24 – 29 , 20 , 30 ] and D.

meteloides [

20 , 31 ]. However, daturas are far from the economic

importance for global agriculture that other Solanaceae have

( tomato , potato , pepper , etc.). This is the reason why the number

of articles and discoveries produced in the last decades using daturas

as experimental system is limited. At present, the trend in DH

research is to use model species (rapeseed, barley , tobacco , etc.) to

study fundamental aspects of the process, and to use recalcitrant

crops to try to make them responsive.

Androgenesis in Solanaceae

214

3 Tobacco

After more than 500 years of cultivation, tobacco ( Nicotiana taba-

cum ) is considered the most valuable non-food crop in the world.

Among the 178 primary crops listed in the 2012 FAOSTAT data-

base [

32 ], tobacco ranks 49th in area harvested (4,291,014 Ha)

and 82nd in production, with 7,490,661 t. Its main utility is the

production of cigars, cigarettes and other derivatives used by the

tobacco industry. Nowadays, the health problems associated to the

habit of smoking are causing a decrease in the traditional uses of

tobacco. However, tobacco is especially suitable for genetic trans-

formation, which makes this crop a good candidate to be exploited

as a biofactory. Indeed, tobacco can be used to produce starch for

bioethanol or for industrial purposes [

33 , 34 ], to produce vaccines

[

35 – 37 ], and a large list of other pharmaceuticals [ 38 – 41 ].

Few years after Guha and Maheshwari milestone report, sev-

eral groups published in a time range of 2 years the production of

haploid plants from tobacco anthers [

42 – 45 ]. A representative

example was the work of Nitsch and Nitsch [

42 ], who published

on Science a paper presenting a method “ by which hundreds of

haploid plants of various species of Nicotiana can be raised from

pollen grains ” . Soon after them, several researchers studied this

phenomenon in tobacco from different experimental approaches

[

46 – 53 ]. In addition to N. tabacum , pioneering researchers also

explored and successfully achieved the induction of haploidy in

other Nicotiana species, including N. sylvestris , N. afﬁ nis , N. rus-

tica , N. attenuata , N. knightiana , and N. raimondii [

42 , 54 ].

Since then, tobacco has been considered for long as a model spe-

cies where to induce microspore embryogenesis efﬁ ciently. Indeed,

there are different well set up protocols currently available to

obtain DHs from anther and isolated microspore culture s with an

acceptable efﬁ ciency [

55 – 58 ]. For anther culture , most of the pro-

tocols include a cold treatment of excised ﬂ ower buds prior to

anther excision and culture on a charcoal-containing medium

[

55 ]. However, its relative simplicity makes isolated microspore

culture the method of choice. For microspore culture, the most

common way to stress the microspores is to starve them from car-

bon and nitrogen sources while applying a inductive, mild heat

shock [

55 , 56 , 59 ]. After induction, embryogenic microspores are

transferred to a carbon and nitrogen-containing medium where

they continue dividing and grow into haploid embryos. These

embryos are transferred to a low- sucrose , agar -based solid medium

for germination , and are ﬁ nally treated with colchicine solutions

for genome doubling [

55 ]. Aside of these standard methods, the

ﬂ exibility of tobacco microspores allowed the application of differ-

ent types of stresses to induce the androgenic switch. Although

these are not the most efﬁ cient ways to induce tobacco micro-

spores, successful induction to embryogenesis has been achieved,

Jose M. Seguí-Simarro

215

for example, by the application of basic pH (8–8.5), lithium (5 mM

LiNO

3 ), abscissic acid (0.01 mM), reduced atmospheric pressure

(12 mmHg) or centrifugation at 10,000–11,000 g (reviewed in

ref. [

60 ]). As in other model species like rapeseed, tobacco micro-

spores also offer the possibility to reproduce microgametogenesis

in vitro, provided that microspores are cultured in a rich, non-

starving medium, with no stress sources [

61 , 62 ]. In this way,

mature and fertile tobacco pollen can also be obtained in a petri

dish. Thanks to these well- established protocols, tobacco embryo-

genic cultures and microspore- derived DH plants can now be rou-

tinely generated, and serve as excellent tools for the study of many

different basic and applied aspects of the process of microspore

induction and embryo development . In the last 15 years, the

majority of the studies published on tobacco microspore embryo-

genesis used this species as a model system where to study the

changes undergone by the induced microspore at the physiologic,

transcriptional, metabolic, or ultrastructural levels, among others.

In particular, tobacco microspore embryogenesis was used to deci-

pher the cellular and ultrastructural changes undergone by the

induced microspore [

20 , 48 , 49 , 51 , 63 , 64 ] and speciﬁ cally by

plastids [

20 ], as well as to discover speciﬁ c mRNAs [ 65 , 66 ], MAP

kinases [

67 ], phosphorylated proteins [ 68 – 70 ], metabolites [ 66 ],

and heat shock gene expression [

71 ] associated with the induction.

These studies have contributed signiﬁ cant insights in the under-

standing of how and why microspores are reprogrammed towards

embryogenesis.

Aside from basic studies, tobacco has also been used to explore

the advantages of DH technology in plant breeding. In addition to

the production of pure lines for hybrid seed production, another

advantage of DH technology is the avoidance of hemizygous trans-

formants when combined with genetic transformation. Such a

combination of both technologies was used in 2007 to produce an

innovative breeding technology. Ribarits et al. [

72 ] produced

reversible male-sterile tobacco plants by ﬁ rstly introducing mutated

tobacco glutamine synthetase genes fused to the tapetum-speciﬁ c

TA29 and the microspore -speciﬁ c NTM19 promoters, and sec-

ondly producing a non-segregant, male-sterile DH population

through microspore culture . In this population, male sterility could

be overcome at will by the exogenous addition of glutamine to

plants or to in vitro maturing pollen . This is an interesting example

of how this technology can help in plant breeding beyond the mere

production of DH pure lines.

In conclusion, tobacco has served during many years as a useful

model system to advance in the knowledge of microspore embryo-

genesis . In the last years, it appears that its role as a prominent dicot

model has been taken by rapeseed ( Brassica napus ), which is the

dicot model species used for most of the recent studies at the cel-

lular, molecular and genetic levels. From the applied point of view,

Androgenesis in Solanaceae

216

the decrease in tobacco consumption worldwide may have an

impact on tobacco breeding programs and therefore on the use of

tobacco DHs in such programs. However, the suitability of tobacco

for both genetic transformation and microspore embryogenesis

could open the door for a promising future of doubled haploid y in

this crop as a tool to be combined with transformation.

4 Potato

Potato ( Solanum tuberosum ) is a solanaceous crop originally from

South America. It is believed that the ﬁ rst places where potato was

cultivated were the region of the Titicaca Lake, in the north of

Bolivia, and the highlands of the Andes [

73 ]. Andian populations

of the north of Peru and the south of Bolivia were the ﬁ rst to eat

wild potatoes around 3000–4000 years bc . It was introduced to

Europe by Spanish expeditions through Seville in 1570, and later

on it was extended to the rest of Europe [

73 ]. The edible part of

potato plants are their tubers, which are nowadays extended world-

wide as an essential part of many cuisines, as well as of a wealth of

processed foods. Indeed, potato ranked eighth in production

(365,365,367 t) and 18th in area harvested (19,278,549 Ha)

among the 178 different crops analyzed in the 2012 FAOSTAT

database [

32 ]. These data illustrate to what extent potato is impor-

tant in the current world’s economy.

The major cultivated species of potato ( S. tuberosum ssp.

tuberosum ) is an autotetraploid. Other interesting potatoes include

cultivated species like the tetraploid S. tuberosum ssp. andigena , the

diploid S. stenotomum and S. phureja , and the diploid wild potato

relative S. chacoense . These species are principally used for genetic

studies and for plant breeding, for example as sources of resistance

for certain diseases. Despite that potato is a sexual species, it is often

difﬁ cult to have it sexually reproduced. Sexual crosses are usually

restricted to breeding centers which use them to generate new vari-

eties, taking advantage of their germplasm collections from differ-

ent potatoes and related species [

73 ]. However, cultivated potatoes

are vegetatively propagated, using the adventitious buds formed on

the tubers. In this way, propagation is easier and the populations

obtained are homogeneous. However, this has generated a very low

level of genetic variability among the different potato cultivars. This

is why it is highly desirable to obtain reduced, dihaploid and even

monohaploid plants for potato breeding at the haploid level and for

genetic analysis [

74 – 76 ]. For example, genetically heterozygous

potato dihaploids may be used as parents to hybridize with other

Solanum species in order to obtain diploid, tetraploid, or even

higher ploidy individuals with new genetic combinations.

Sometimes, these hybrids are sterile and haploidization is employed

as a possible strategy to overcome reproductive barriers, as recently

Jose M. Seguí-Simarro

217

reported in hybrids between Solanum bulbocastanum and S.

tuberosum [

77 ]. Examples on the use of androgenic approaches in

potato breeding also include S. chacoense x S. phureja clones [

78 ]

and S. brevidens x S. tuberosum somatic hybrid s [

79 ]. Other exam-

ples of this and other approaches to obtain potatoes with new

genetic backgrounds can be found in a review by Tai [

80 ].

Different techniques for dihaploidization have been described

in the literature. These include interspeciﬁ c hybridization of culti-

vated potato and related Solanum species with certain haploid

inducer clones of S. phureja , and microspore embryogenesis

(reviewed in refs. [

81 , 82 ]). With respect to the latter, potato can-

not be considered as a model system, but at present there are sev-

eral methods capable to induce androgenesis from anther culture s

[

80 , 83 , 84 ]. The main difference between them consists on the

use of different culture media, either liquid [

85 – 87 ] or semisolid

[

79 ]. In a technical review of 2003, Rokka [ 84 ] described a method

for potato anther culture based on the Murashige and Skoog (MS)

basal medium [

88 ], capable of inducing microspore embryogene-

sis in a wide range of genetically diverse potato species, including

interspeciﬁ c and intergeneric hybrids between them. Aside of S.

tuberosum , other papers have reported induction of androgenesis

by anther cultures of S. phureja [

89 – 93 ], and other potato relatives

such as S. acaule [

94 ], or S. chacoense [ 95 – 98 ].

The effect of colchicine in anther culture s was also evaluated

[

89 ], concluding that it did not affected neither the efﬁ ciency of

induction nor the percentage of monoploid plants obtained.

Anther culture s have been described to produce microspore-

derived embryos and calli. Direct embryogenesis is preferred over

callus formation due to the lower occurrence of somaclonal varia-

tion [

82 ]. However, a RAPD analysis of the plants obtained from

S. phureja anther cultures revealed the occurrence of genetic

clones, possible originated by secondary embryogenesis during

anther culture [

89 ]. Other studies, also based on molecular mark-

ers, have revealed interesting results. For example, Sharma et al.

[

99 ] used SSRs to demonstrate that anther culture-mediated

dihaploidization of S. tuberosum tetraploids involves extensive

changes and genetic rearrangements. In addition, they demon-

strated the occurrence of somatic embryo genesis from anther

walls, and of somaclonal variation in the tetraploid somatic regen-

erants. Birhman et al. [

97 ] studied by means of RFLPs the genetic

architecture and the origin of S. chacoense plants produced through

anther culture. They showed that some of the plants obtained had

different ploidy levels but their genetic constitution was identical,

which suggested the occurrence of microspore-derived, mixoploid

calli developing clonal plants from their ploidy-different cells. They

also showed that some of their plants, although regenerated from

the same callus, had different genetic constitutions, which led

them to conclude that they might come from two microcalli

Androgenesis in Solanaceae

218

derived from two different microspores, but proliferating together

within the anther.

Although not abundant, reports exist on potato isolated micro-

spore culture s, too [

100 – 103 ]. Through the application of cold and

starvation treatments, one of these reports [

103 ] described a per-

formance better than anther culture s and, as expected, a depen-

dence on the genotype of the donor plant. They also observed that

microspores of a dihaploid genotype divided symmetrically after 3

days from isolation, giving rise to suspensorless embryos. Symmetric

divisions and suspensorless MDEs have also been observed in the

rest of reports dealing with potato microspore cultures. However,

microspores of “Albina,” a tetraploid potato, divided later (8 days

after isolation) and asymmetrically, giving rise to a large and a small

cell, which led to the formation of a zygotic-like suspensor and an

embryo proper, respectively [

103 ]. This observation is interesting,

since the number of inducible species that develop suspensor-

bearing MDEs is very limited, being Brassica napus the most prom-

inent example. However, none of the above mentioned reports

have demonstrated the successful production of viable potato plants

from isolated microspore cultures. Therefore, there is still work to

do in order to provide a reliable and complete protocol.

5 Tomato

Within the genus Solanum , the section Lycopersicon includes toma-

to es and their wild relative species. The native distribution of this

section ranges ac ross the west-central part of South America, from

the high Andes to coastal Ecuador, Peru and the north of Chile,

although it seems that it was in Mexico where tomato was domes-

ticated, and then introduced to Europe and Asia by the Spanish

and Portuguese [

73 ]. Although the taxonomy of the section

Lycopersicon has been subjected to a long-lasting discussion that

has not yet reached a widely accepted consensus, the most recent

classiﬁ cation [

104 ] divided the section into four groups: the

Lycopersicon group ( S. lycopersicum , S. pimpinellifolium , S. chees-

maniae , and S. galapagense ), the Neolycopersicon group ( S. pen-

nellii ), the Eriopersicon group ( S. peruvianum , S. corneliomulleri ,

S. huaylasense , S. habrochaites , and S. chilense ), and the Arcanum

group ( S. arcanum , S. chmielewskii , and S. neorickii ). From an

economic point of view, cultivated tomato ( S. lycopersicum ) is by

far the most important of the section. Tomato is the ﬁ rst vegetable

crop worldwide, both in terms of production (161,793,834 t in

2012) and cultivated area (4,803,680 Ha in 2012). Among the

178 different crops analyzed in the 2012 FAOSTAT database [

32 ],

tomato ranks 16th in production and 47th in area harvested. These

data give an idea of the tremendous importance that tomato has

for global agriculture.

Jose M. Seguí-Simarro

219

However, and despite of its importance, little is known in the

tomato DH ﬁ eld, with no reliable and standardized methods avail-

able so far. Over the past 40 years, a signiﬁ cant number of media

types and conditions, as well as combinations of nutrients, vita-

mins, growth factors and supplements, have been assessed

(reviewed in ref. [

105 ]). During these years most of the published

papers reported just on induction of calli [

106 , 107 ] or multicel-

lular structures [

108 ], or regeneration of roots [ 109 ] or apical

shoots [

110 ]. Only two laboratories have published the complete

regeneration of entire tomato plants with a demonstrated haploid

or DH origin [

111 – 115 ], although with different levels of mor-

phological variability ranging from high [

116 ] to low [ 114 ]. In all

of these published studies, mixoploidy in a percentage of individu-

als and a low general efﬁ ciency were common features [

114 , 115 ].

Aside of these reports, a third group reported in 2001 the produc-

tion of androgenic plants, but not in tomato. Gavrilenko et al.

[

117 ] obtained plants by culturing anthers of a somatic hybrid

between S. lycopersicum and S. etuberosum , a wild non-tuberous

Solanum species with several desirable agronomic traits. Again, the

androgenic efﬁ ciency obtained from this intergeneric hybrid was

extremely low, with 3.4 % of responding anthers and ﬁ ve plants

from one of the hybrid donor plants, and 1.2 % of responding

anthers and only one plant from the other hybrid donor plant

used. The embryogenic response of anther culture s of S. lycopersi-

cum x S. peruvianum hybrids was tested by Cappadocia and

Ramulu [

118 ]. Although they reported the observation of the ﬁ rst

stages of embryogenesis, the plants obtained derived from anther

tissues. Out of the S. lycopersicum species, anther cultures have also

been performed in S. peruvianum [

119 , 120 ]. Reynolds [ 121 ]

reported the production of callus, embryoids and regenerated

plants from cultured anthers of wild tomato ( S. carolinense ), a pas-

ture weed from North America. In this work, the occurrence of

microspore- derived callus or embryoids depended on the hor-

monal composition of the culture medium. Other reports on S.

carolinense anther culture can be found in refs. [

122 – 126 ]. It can

be deduced from all these works results that the state of the art on

DH research in tomato is far from being considered minimally

useful to be applied for DH production on a routine basis. Indeed,

tomato can be regarded as one of the major examples of species

recalcitrant to androgenesis induction. More efforts are needed to

obtain successful results.

As in other in vitro morphogenic processes, the most critical

factors to obtain androgenic DHs from tomato are the genotype

and the developmental stage. As for the genotype, male-sterile

mutant lines have been shown to be especially sensitive to being

induced [

112 , 114 , 115 , 127 , 128 ]. Usually, male-sterility is asso-

ciated to defects at the late meiocyte stage, which ends up degen-

erating and dying. This was demonstrated for the ms10

35

Androgenesis in Solanaceae

220

male-sterile genotype, where most of the work on tomato DHs has

been developed [

115 ]. Aside of male-sterile mutants, all of the

works published to date on the evaluation of androgenic compe-

tence in commercial tomato cultivars have reported null or very

few positive results. It is important to highlight that this informa-

tion comes from published papers, but it might not be the only

one. Considering the extraordinary importance of this crop for

breeding companies, some of them have set up speciﬁ c research

programs and collaborations to ﬁ nd the key switch for this species.

Indeed, some methods claiming the development of protocols to

obtain haploids in tomato have been patented [

5 , 129 , 130 ]. Thus,

it might be possible that some companies would have found a way

to obtain androgenic DHs in their particular tomato cultivars, but

it is unlikely that this proprietary information will see the public

light in the short-mid-term. The second key factor is the identiﬁ ca-

tion of the right stage for the tomato gametophyte (or its precur-

sor) to be reprogrammed. This has been a matter of debate along

40 years of research on tomato DHs. A number of works have

been sporadically published [

109 , 111 , 127 , 131 – 135 ], but a clear

consensus could not be obtained from them. Perhaps, the most

suggestive work of the ﬁ rst years of tomato haploidy research was

by Dao and Shamina [

135 ]. They induced the formation of callus-

like structures from anthers with meiocytes at the tetrad stage, just

after walling. On the other hand, they also obtained embryoids

directly from anthers containing late, vacuolate microspores or

young, bicellular pollen grains. This work, not further continued,

pointed to the notion that, as also suggested by Chlyah et al. [

136 ],

tomato could follow the trend observed in other species, where

different stages may be responsive under different treatments

[

137 – 139 ]. In the last decade, several studies have reinforced this

idea. On the one hand, the optimal stage for anther culture was

narrowed to the meiocyte just before compartmentalization of the

tetrad [

111 , 113 ], which is a quite infrequent feature of tomato,

together with some species of the genus Vitis [

140 ]. This develop-

mental window implies that recombination must be successfully

ﬁ nished without disruption, but microspore formation (tetrad

compartmentalization) has to be prevented. On the other hand,

the formation of few-celled structures from isolated microspore

culture s was described upon exposure to a combined treatment

of starvation, cold and colchicine [

108 ]. In 2007, it was ﬁ nally

demonstrated that in tomato, gametophyte precursors can be

induced to divide at two different stages: meiocytes and vacuolated

microspores [

114 ]. However, both possibilities still have a great

number of limitations, as explained in the next paragraphs.

Haploid and DH plants can be obtained from in vitro cultured

anthers containing meiocytes (Fig.

1 ), but they are not the only

products of this process. Plants originate from meiocyte-derived

calli, which may in turn come from two different origins: (1) from

Jose M. Seguí-Simarro

221

haploid meiotic products still enclosed within the tetrad (the future

microspore ), that stop their gametophytic program and start pro-

liferation, or (2) from proliferating diploid cells, produced from

the fusion of two different meiotic products. As for the ﬁ rst origin,

it was demonstrated that stress-induced meiocytes (Fig.

1a )

undergo a series of defects in tetrad compartmentalization (forma-

tion of defective, incomplete, or absent cell wall s) that facilitate the

fusion of nuclei between adjacent, not well separated cells [

114 ,

115 ], in a way similar to that described for the tomato mutant

Fig. 1 Tomato anther and microspore culture . ( a ) Meiocytes within a cultured

anther. Arrowheads point to two nuclei coalescing in the same cytoplasm. ( b )

Callus ( arrowhead ) emerging from a cultured anther. ( c ) Cultured callus with

shoot- forming organogenic buds. ( d ) Plantlet regenerating from a callus. ( e )

Haploid and doubled haploid tomato plants. ( f , f′ ) Isolated microspore culture

( arrowheads point to the nuclei of a dividing microspore). ex exine. Bars: a , 20

μm; b , 2 mm; c , d , 1 cm; f , f′ , 15 μm Images adapted from [ 114 ] .

Androgenesis in Solanaceae

222

pmcd1 ( pre-meiotic cytokinesis defect 1 ), which generates diploid

gametes due to the fusion of two haploid meiotic products before

microspore release [

141 ]. When callus proliferation starts ﬁ rst and

nuclear fusion occurs later on, the new cell formed will have a dip-

loid nuclear content, formed by two identical haploid genomes. In

other words, it will be a DH cell that can potentially give rise to a

DH callus (Fig.

1b–d ) and a DH plant (Fig. 1e ). Alternatively,

nuclear fusion may occur late in callus proliferation, giving rise to

mixoploid calli where some cells remain haploid, and others,

derived from those undergoing nuclear fusion, will be DH [

114 ,

115 ]. A similar mechanism for mixoploid (haploid + DH) callus

formation was proposed to explain the occurrence of genetically

identical (clonal) plants with different ploidy levels from

microspore- derived calli of S. chacoense [

97 ]. However, nuclear

fusion of haploid tetrad cells may occur ﬁ rst, followed by prolifera-

tion. This will lead to the second of the two possible origins men-

tioned above. In this case, the two fusing nuclei will also be haploid,

but genetically different. Since they come from meiotic recombi-

nation at prophase I, fusion of two reduced meiotic products

would generate new allele combinations not necessarily homozy-

gous . Therefore, the callus and plant coming from this cell would

not be DH. Interestingly, Birhman et al. [

97 ] demonstrated by

RFLP genetic analysis in S. chacoense that plants regenerated from

the same callus may be genetically different. As explained in the

section devoted to potato , they explained this result as the coupled

proliferation of two calli originated from two different microspores.

However, in light of what occurs in tomato, their results might be

alternatively interpreted as the proliferation of a single callus com-

ing from a diploid microspore, whose single diploid nucleus comes

from the fusion, during meiosis, of two different haploid nuclei of

a non-well compartmentalized tetrad. In addition to these two

possible origins, callus and plants may also come from the prolif-

eration of somatic, anther tissues. Indeed, ﬁ lament tissues typically

exhibit a high proliferative response when cultured in vitro [

142 ],

and it is believed that tomato anther tissues at meiotic stages are

more sensitive to tissue culture than those of older stages [

105 ].

All this considered, it seems that DHs are not the only individuals

that can arise from tomato anther culture s. So the question is: how

frequently do anther-derived tomato DHs arise? In a recent study,

it was shown that the ﬁ rst possibility (proliferation and then fusion)

accounts for only 7 % of the plants regenerated, which would be

haploid or DH [

115 ]. This study also revealed that the second pos-

sibility (fusion ﬁ rst and then proliferation) accounted for 10 % of

the plants, and the third possibility (somatic origin) accounted for

83 % of the plants. In other words, more than 90 % of the plants

obtained have a non-haploid origin, and only 7 % would be useful

to obtain DH plants. It is evident that non-DH plants represent

the vast majority of plants produced. It is also evident that this

Jose M. Seguí-Simarro

223

method requires the genetic evaluation of each single regenerant

to determine their origin and ploidy, and that most of them are

useless and should be discarded. In other words, this technology,

at its current state of the art, has few chances to be implemented by

breeding companies to produce DHs at large scale.

Microspore culture is the second option to explore to obtain

androgenic DHs in tomato . When microspores at the vacuolated

stage are isolated and grown in liquid medium, it is possible to

induce proliferative divisions in these microspores (Fig.

1f, f ′), gen-

erating callus-like [

108 ] and embryo-like structures [ 114 ]. This

alternative would prevent all the problems mentioned above as

derived from using anthers and meiocytes. However, this approach

is still at its infancy. Up to now very few genotypes have been

assessed using this method, and in the very few positive results

obtained, it has not been possible to go beyond the ﬁ rst divisions

of the developing MDEs [

114 ]. After these divisions, embryos

arrest and never progress beyond the early globular stage [

114 ].

Perhaps, culture conditions were not well optimized to allow for

in vitro embryo development , with no zygotic endosperm to pro-

vide nutrients and developmental cues to ensure proper embryo

development. It could also be possible that haploidy would unmask

embryo-lethal genes in the dividing haploid cells, which would

preclude further development.

As seen in this section, current research in tomato haploidy is

still far from providing a reliable and efﬁ cient protocol to produce

androgenic DHs. After more than 40 years of tomato DH research,

little is still known about the origin of the recalcitrance shown by

cultivated tomato. In this respect, one interesting hypothesis was

that suggested by Sangwan and Sangwan-Norreel [

20 ], who

related recalcitrance with plastid differentiation. These researchers

studied the ultrastructure of plastids in an extensive number of spe-

cies that, according to their own results, they divided into

androgenic and recalcitrant . They found that in the androgenic

group, proplastids do not differentiate into amyloplasts until the

mid or late bicellular pollen stage of microgametogenesis, whereas

in the recalcitrant group, such a differentiation occurs soon, during

microspore development, or even sooner, before tetrad formation.

In other words, they suggested that the differentiation of proplas-

tids into amyloplasts marks the end of a favorable period for andro-

genesis induction. This has been widely acknowledged later on,

accepting that starch accumulation is indicative of commitment

towards gametogenesis (reviewed in refs. [

143 – 145 ]). Interestingly,

their investigations showed that, as opposed to the rest of the

Solanaceae studied, tomato plastids soon differentiated into amy-

loplast, being present in microspores before the ﬁ rst pollen mitosis.

Thus, the difﬁ culty of induction shown by tomato microspores

might be related to the fact that their plastids accumulate starch

unusually soon, as compared with other, more sensitive species like

Androgenesis in Solanaceae

224

rapeseed, or even within their same genus, tobacco or Datura .

This possibility is also consistent with the wide consensus about

the difﬁ culty of inducing in vitro redifferentiation or organogene-

sis from starch-rich somatic tissues, compared to tissues where

plastids are still in the form of proplastids or chloroplasts. As an

alternative to the androgenic approach, a limited number of works

have been published on the development of ways to haploidy such

as gynogenesis [

117 , 146 ], ovary culture, pollen irradiation [ 147 ],

or wide hybridization, and little positive results have been pub-

lished (reviewed in ref. [

105 ]). Perhaps, the future of tomato DH

research should focus on exploring the possibilities of microspore

culture , and in parallel, of new, alternative pathways to doubled

haploid y not related to androgenesis. An example of alternative

ways to approach tomato DHs might possibly be the one demon-

strated by Ravi and Chan [

148 ] in Arabidopsis : the production of

haploid plants produced by uniparental centromere-mediated

genome elimination. By manipulating a single centromere-speciﬁ c

histone (CENH3), it is possible to produce cenh3 null mutants

expressing altered CENH3 proteins. When crossed to wild type

plants, chromosomes from the mutant are eliminated, thereby pro-

ducing haploid and DH progenies. However, this approach has

still a number of technical challenges still to be overcome in order

to make it possible in tomato.

6 Eggplant

Eggplant ( Solanum melongena L.), also known as aubergine, brin-

jal, or Guinea squash [

149 ], is another of the most important veg-

etables worldwide. In 2012, eggplant ranked the sixth and eighth

among all the vegetable crops for production and area harvested,

respectively [

32 ]. In 2012, 1,853,023 Ha of eggplant ﬁ elds were

cultivated, and a total of 48,424,295 t were produced in the world

[

32 ]. From these, nearly 85 % were produced in China (59.5 %)

and India (25.2 %), the two main producers worldwide. Eggplant

is thought to have its origins in Asian tropical and subtropical

regions, where it extended to Africa and then to the Mediterranean

area of Europe [

73 ]. Eggplant has a remarkable importance in eco-

nomic terms in China, India, Africa, and some subtropical, Central

American countries. It is also grown in some warm, temperate

regions of the south of the USA, and in the Mediterranean basin,

in countries such as Italy, Greece, or Spain, where eggplant consti-

tutes a classic ingredient in the renowned “Mediterranean diet.”

With respect to haploidy induction, eggplant appears to be

between tomato and tobacco in terms of recalcitrance. It is possi-

ble to induce the deviation of the eggplant microspore within the

anther towards an embryogenic route, thus generating a haploid

or DH embryo that will eventually germinate into a haploid or DH

Jose M. Seguí-Simarro

225

plant. However, we still are far away from the efﬁ ciency shown by

tobacco. The ﬁ rst report of plant regeneration from anther cul-

ture s dates from 1973 [

150 ]. Here, authors were able to induce

callus proliferation and emergence from anthers, and plant regen-

eration upon treatment with colchicine . However, according to

the authors’ own interpretation, it is likely that the generated calli

were produced from the connective tissue of the anther, having

therefore a non haploid origin. Few years later, other articles

reported the production of eggplant haploid individuals [

151 ,

152 ], or the induction of callus derived from microsporogenous

tissue within the eggplant anthers [

153 ]. From these calli, they

were able to regenerate shoots and roots. Then, Robert Dumas de

Vaulx and Daniel Chambonnet established the basis for a general,

reliable and reproducible protocol for haploid embryo and DH

production from eggplant anther cultures [

154 , 155 ]. This

method, in its original formulation, consisted on the incubation of

anthers in a medium containing 2,4-dichlorophenoxyacetic acid

(2,4-D) and kinetin, at 35 °C in darkness during 8 days to induce

embryogenesis. Then, cultures are moved to 25 °C and exposed to

light in order to promote the development of MDEs within the

cultured anthers. At day 12th, anthers are moved to a second

medium with a reduced level of 2,4-D and kinetin. This has been

the basis of many protocols, now routine methods, adapted to par-

ticular eggplant varieties that have been, or are currently, used in

breeding programs. At present, pure DH lines of some varieties

and hybrids have already been developed [

156 – 159 ], based on

modiﬁ ed versions of this protocol. As it happens in all other induc-

ible species, the efﬁ ciency of the embryogenic response of the

microspores in eggplant is greatly inﬂ uenced by the genotype of

the donor plant [

159 ] and by the stage of microspore development

when anthers are excised. Related to the latter, a recent report

highlighted two particular features of eggplant that may have a

signiﬁ cant impact on the efﬁ ciency of induction. The ﬁ rst is the

particular thickness of eggplant anther walls, which seems to delay

the access of inductive factors to the anther locule, thus reducing

their effect over inducible microspores [

158 ]. Therefore, the cul-

ture of anthers with younger microspores was proposed to allow

for younger microspores to grow up to the inducible stages while

factors are entering the locule. The second is the heterostyly, pres-

ent in certain eggplant cultivars, which might have an inﬂ uence in

the embryogenic response. Salas et al. [

158 ] studied the embryo-

genic response of short and long-styled buds present in a hetero-

stylic cultivar, and demonstrated that each ﬂ oral morph produced

buds and anthers of different lengths, but equally useful for anther

culture, since both morphs produced similar embryogenic

responses. Microspore embryogenesis depends on culture

conditions, too, including temperature, type, and concentration of

growth regulators [

156 , 157 , 160 , 161 ]. Genetic variability was

Androgenesis in Solanaceae

226

also observed in plants regenerated from anther cultures [ 162 ].

These results conﬁ rmed that in general, eggplant microspore

embryogenesis in cultured anthers behaves as in other better-

studied species, although with some particularities inherent to this

species. Overall, eggplant anther cultures can be and are being

used to obtain DHs for breeding purposes [

156 – 159 , 163 – 165 ].

Two examples can illustrate the potential of this technology in egg-

plant. First, in 2002 Rizza et al. [

166 ] generated a population of

androgenic dihaploid plants, derived from somatic hybrid s between

S. melongena and a wild eggplant relative, S. aethiopicum , and

demonstrated that they constitute a useful source for the introduc-

tion into cultivated eggplant varieties of the Fusarium oxysporum

resistance typical of S. aethiopicum group Gilo. Second, a similar

approach was used to reduce to the dihaploid status the ploidy of

eggplant somatic hybrids between S. melongena and other wild

relative, S. integrifolium , in order to facilitate their crossability with

cultivated eggplant varieties for the introgression of Fusarium

resistance [

167 ].

Despite that eggplant embryos can be successfully induced

from microspores cultured within the anther, the development of

a method for embryogenesis induction from isolated eggplant

microspores would be highly desirable. Although the occurrence

of somatic regenerants derived from anther walls seems not to be a

big issue in eggplant [

159 ], isolated microspore culture would

avoid the other problems mentioned in the introduction: the

uncontrolled contribution of tapetal cells to culture conditions and

the low efﬁ ciency. Related to this, it is striking that very few papers

have been published on the successful production of DH plants

from isolated microspores [

168 – 170 ]. Miyoshi described the pro-

liferation of eggplant microspores to form calli, from which he

obtained DHs through organogenesis. He obtained 20–65 calli

per anther, and 0.001–0.02 plantlets were regenerated from each

callus. As mentioned by Miyoshi, this efﬁ ciency was clearly beyond

that of anther culture s. In the last 2 years, two papers have contrib-

uted to the progress in this ﬁ eld, conﬁ rming the applicability of the

studies mentioned above to different eggplant genotypes, and

developing a protocol that further enhances the efﬁ ciency of micro-

spore induction [

169 , 170 ], well above that previously published

by Miyoshi [

168 ]. Like in the above mentioned paper, the plants

apparently did not come from embryos, but were regenerated

through a callus phase instead. However, a careful study of the

process of microspore proliferation showed that, actually, micro-

spore embryogenesis seems to initiate as in other inducible species

(Fig.

2a–c ), but arrests at the globular embryo stage (Fig. 2d ).

Instead of experiencing the radial-to-bilateral symmetry transition

typical of zygotic embryo s, eggplant MDEs enter a proliferative

phase of undifferentiated growth and become callus-like structures

(Fig.

2e ) [ 169 ]. Haploid and DH plants regenerate from these calli

Jose M. Seguí-Simarro

227

Fig. 2 Microspore culture s in eggplant . ( a , a′ ) Two-celled, induced microspore ( arrow ) together with unicellular,

non-induced microspores ( arrowheads ). Note the blue , DAPI -stained nuclei in a′ . ( b , b′ ) Four-celled, dividing micro-

spore ( arrow ). ( c , c′ ) Multicellular microspore. ( d , d′ ) Globular embryo. ( e ) Embryo-derived calli ( a – d are phase

contrast images, a′ – d′ are ﬂ uorescence images of DAPI-stained samples). Bars: a – d , a′ – d′ , 50 μm; e , 1 mm

through organogenesis, as ﬁ rst described by Miyoshi. In order to

further increase the efﬁ ciency of induction, but principally to avoid

the transformation of MDEs into calli, a number of experimental

factors were tested, including polyethylene glycol ( PEG ), manni-

tol , abscisic acid, epibrassinolide, naphthaleneacetic acid (NAA),

6-benzylaminopurine (BAP), arabinogalactan protein s ( AGPs ),

and different combinations of them [

169 , 170 ]. It was found that

Androgenesis in Solanaceae

228

certain combinations of these factors increased the efﬁ ciency of

microspore induction towards embryogenesis, but only one

(AGPs) permitted the development of bipolar embryos [

170 ].

These embryos exhibited anatomically normal hypocotyls and

radicular poles. However, their shoot apical meristems and cotyle-

dons were either absent or severely distorted, which precluded a

normal germination . Together, these results extended the knowl-

edge, still scarce, regarding microspore culture in eggplant. In

addition, they clearly pointed to deﬁ ciencies in the composition of

the culture medium where the post-inductive phases take place, as

the responsible of this abnormal pattern of embryogenesis. Thus,

further studies to optimize eggplant microspore cultures should

focus principally on the elimination of the bottleneck by which

eggplant MDEs are not capable of progressing beyond the globu-

lar stage to produce mature, well-formed embryos ready for germi-

nation. In this way, the phase of plant regeneration from calli

through organogenesis would be eliminated, and the method

would be considerably faster, cheaper, and, therefore, efﬁ cient.

7 Pepper

Peppers (genus Capsicum ) are plants native of America, where they

were cultivated for thousands of years by the people of the tropical

regions of South America [

73 , 171 ]. At present, they are cultivated

worldwide. The genus Capsicum consists of approximately 20–27

species, ﬁ ve of which are domesticated: C. annuum , C. baccatum ,

C. chinense , C. frutescens , and C. pubescens [

171 ]. The fruit of

Capsicum plants has a variety of names, depending on the region

and the type of pepper , including chili pepper, red or green pepper,

bell pepper, sweet pepper, just “ capsicum ” in Australian and Indian

English, or paprika in Hungary. Some of the members of Capsicum

are used in fresh as vegetables, and others are dried and used as

spices. With a total world production in 2012 of 31,171,567 t in

1,914,685 Ha, peppers ranked 44nd in terms of production and

69th in terms of area harvested among the 178 primary crops listed

in the FAOSTAT 2012 database [

32 ]. China was by far the leader

in production of freshly edible peppers with 16,000,000 t, around

51 % of the total production [

32 ]. In dry pepper, India was the

world leader with 1,299,940 t, almost 39 % of the total world

production.

Pepper is the third solanaceous crop that could be deﬁ ned as

difﬁ cult with respect to the induction of androgenesis . In this spe-

cies, the production of haploids and DHs has been assessed by dif-

ferent means including the use of both the male and the female

gametophyte and their precursors (reviewed in ref. [

172 ]). Apart

from the spontaneous occurrence of some cases of in vivo andro-

genesis of no practical relevance [

173 ], haploids in pepper were

Jose M. Seguí-Simarro

229

ﬁ rstly obtained through parthenogenesis (reviewed in refs. [ 172 ,

174 ]). However, after the discovery of in vitro anther culture as a

way to induce androgenesis [

1 ], this has been the most successful

technique so far. The ﬁ rst reports on the production of haploid-

Capsicum plants by anther culture involved the use of Asian variet-

ies [

175 – 177 ]. A year after, anther culture of European varieties

was reported [

178 , 179 ]. Thereafter, many C. annuum cultivars

and interspeciﬁ c crosses have been tested for responsiveness to

anther culture [

180 – 190 ]. To our knowledge, the most extensive

of this type of studies was that of Gémes-Juhász et al. [

191 ], who

tested over 2000 genotypes, including Hungarian sweet, Hungarian

spice, Dutch blocky, Spanish, and Turkish pepper types. Anther cul-

ture s have been used as a tool for genetic- [

192 ] and cell biology-

based studies [

193 – 201 ], as well as for dissecting the genetic basis

of resistance to pests [

202 ] and diseases [ 203 ]. However, the most

important application of anther-derived, androgenic cultures in

pepper has been their use in breeding programs, ﬁ nally aimed to

produce commercial hybrids with maximum heterosis and new or

improved traits [

172 , 204 – 216 ]. Although the vast majority of the

studies mentioned above have been conducted in C. annuum ,

anther culture has also been explored in other domesticated species,

such as C. frutescens [

217 , 218 ].

As for many other androgenic systems, microspore embryo-

genesis in pepper strongly depends on three critical factors. The

ﬁ rst relates to the donor plants, their endogenous and exogenous

conditions, their growth environment (photoperiod, temperature,

fertilization , irrigation, use of pesticides, age, season), and princi-

pally the genotype, as it occurs in all other responsive species.

However, it must be mentioned that for the particular case of some

pepper cultivars, it is necessary to use antibiotics to prevent con-

tamination due to the presence of endogenous bacteria in ﬂ ower

buds, extremely difﬁ cult to eradicate by the conventional bud sur-

face sterilization. This has also been observed by other researchers

in pepper [

219 ] as well as in other species [ 220 ], which led to the

routine addition of cefotaxime to prevent bacterial growth. The

second critical factor is the optimal microspore stage to apply the

inductive treatment. In pepper, the discrepancies about the optimal

stage have never been as remarkable as in tomato . The literature

shows examples of papers claiming that, for certain genotypes, the

most inducible stage is the vacuolated microspore [

199 ], whereas

others support the early bicellular pollen as the best stage [

193 ]. In

general, the most accepted notion is that the inducible stages

revolve around the ﬁ rst pollen mitosis, as usual for other species.

Perhaps, the fact of deﬁ ning one stage as the most suitable largely

relies on the precision and correctness of the method used to select

anthers with microspores at the right stage. A recent paper describes

a comparison of four of these methods, including morphological

markers such as bud length, anther length, anther pigmentation,

Androgenesis in Solanaceae

230

and calyx/bud ratio [ 221 ]. This work proposes a combination of

calyx/bud ratio and anther pigmentation (once the bud is open

under the ﬂ ow hood), as the most convenient, fast and accurate

way to identify anthers containing vacuolated microspores and

young bicellular pollen. The third critical factor is the culture envi-

ronment, including the stress used for induction (reviewed in ref.

[

174 ]). As for many other androgenic systems, the ﬁ rst protocols

used to culture pepper anthers generated callus, from which plants

were regenerated through organogenesis [

175 – 177 ]. However,

further reﬁ nement of the experimental procedures allowed the

group of Dumas de Vaulx to obtain the direct induction of embryo-

genesis without an intervening callus phase. This, in turn, permit-

ted a signiﬁ cant improvement in the efﬁ ciency of the process of

DH production [

180 , 222 ]. The improvements introduced by this

method were principally based on the use of a high temperature

treatment (35 °C) to induce microspore divisions, and two differ-

ent medium compositions and growth conditions for induction

and embryo development , in a way similar to that developed by the

same group for eggplant . Thus, a general, reliable method for

anther culture in pepper was established. Thereafter, this general

protocol has been applied, with slight modiﬁ cations, to many dif-

ferent pepper varieties. For example, it was recently shown that

different durations (4, 8, 12, and 16 days) of the 35 °C heat shock

applied to different commercial F1 hybrids of the Lamuyo and

California types had signiﬁ cant effects in embryo production, but

also in callus generation, which increased with prolonged expo-

sures in a genotype-dependent manner [

190 ]. Nowadays, these

and other particular adaptations of the general protocol are still

being used to generate DHs for breeding purposes in a number of

varieties [

174 , 184 , 189 , 194 , 196 , 206 , 217 , 223 – 225 ]. Similarly,

several hybrid seed companies use this technique to obtain pure

lines worldwide.

Few years after the Dumas de Vaulx method was developed,

the addition to the culture medium of activated charcoal was pro-

posed to adsorb toxic, metabolism-derived compounds [

223 ,

226 ]. Based on the work of Morrison et al. [ 182 ], Dolcet-Sanjuan

et al. [

227 ] proposed a biphasic medium consisting of a liquid

medium phase poured over a semisolid, agar -based phase with acti-

vated charcoal, and added a signiﬁ cant improvement with the

implementation of a carbon dioxide environment. This method

allowed for the production of embryos of varieties that did not

respond to the method of Dumas de Vaulx, and increased the efﬁ -

ciency of other, poorly responding peppers. However, the techni-

cal difﬁ culty of having a growth chamber with a carbon dioxide

supply has precluded many laboratories from adopting this method

on a routine basis. At present, the simplicity of the Dumas de Vaulx

method makes it the most popular and routinely used.

Notwithstanding this, the anther culture technique also carries the

Jose M. Seguí-Simarro

231

drawbacks described in the introduction, affecting other Solanaceae

such as eggplant or tomato . Thus, in the case of pepper it would be

equally desirable to have a protocol for isolated microspore cul-

ture . This method, although technically much more complex than

anther culture, would provide signiﬁ cant advantages: a higher efﬁ -

ciency, the avoidance of the uncontrollable secretory effect of the

tapetum surrounding the pollen sac of the anther, and especially

the possibility of occurrence of calli/embryos derived from sporo-

phytic tissues. In the last decades, several research groups have

explored the isolated microspore culture pathway. Although many

of them described the formation of MDEs [

174 , 195 , 196 , 199 ],

in most cases embryo development beyond the globular embryo

stage could not be promoted. One exception to this rule was the

work of Supena et al. [

187 , 188 ], who were able to regenerate

haploid plants from microspores isolated from the anther by non-

mechanical means. The technique described by Supena et al. con-

sisted of the preparation of a biphasic medium, different from that

of Dolcet-Sanjuan et al. [

227 ], and the inoculation of anthers

ﬂ oating on the surface of the liquid phase. Under the right condi-

tions, a few days after inoculation the anthers enter into dehis-

cence, open and pour their microspore contents into the liquid

medium. Embryogenic microspores sink and accumulate at the

bottom of the plate, over the semisolid phase. Thus, microspores

are isolated from the anther and allowed to form embryos out of

the inﬂ uence of anther tissues. The main problem of this method,

as admitted by the authors, was the low percentage (20 %) of

normal- looking embryos. However, in 2011 they published a

reﬁ nement of their own protocol whereby the modiﬁ cation of

several physical and chemical parameters raised the percentage of

normal- looking embryos beyond 50 % [

228 ]. The shed-

microspore method has been tested in many different varieties of

Indonesian hot pepper and worked in all of them with different

efﬁ ciencies, much higher to those obtained by other culture tech-

niques. Indeed, the shed-microspore method was tested against

the most popular methods for anther culture, including that of

Dumas de Vaulx and of Dolcet-Sanjuan [

188 ]. The shed micro-

spore method proved to be better that the other two. Recently, it

was applied to some sweet pepper types. More exactly, the perfor-

mances in terms of callus and MDE production of the shed-

microspore method and of the Dumas de Vaulx method were

evaluated in four sweet pepper cultivars [

190 ]. For all genotypes

tested, the protocol of Dumas de Vaulx promoted the induction

and development of MDEs, but also the growth of callus derived

from anther walls. Instead, the shed-microspore method pro-

duced no callus but only embryos. However, the embryo responses

of the cultivars to each treatment was strikingly different, indicat-

ing that there seems not to be a universally useful method to

induce androgenesis in pepper.

Androgenesis in Solanaceae

232

Aside of the shed microspore culture for microspore isolation,

the mechanical isolation of microspores has also been assessed.

Supena et al. [

187 ] reported a high rate of microspore induction in

the same Indonesian hot types evaluated for the shed microspore

method. However, a very low percentage of microspores trans-

formed into MDEs, yielding a maximum of 0.1 regenerated plants

per bud. Higher efﬁ ciencies were obtained in hot pepper types by

Kim et al. [

229 ], using a different protocol deﬁ ned by a pretreat-

ment of microspores at 32 °C in sucrose -free medium, the use of

sucrose as a carbon source for embryo growth, and the use of an

optimal microspore plating density. However, the quality of the

embryos obtained still needed to be improved. Very recently, the

same group published a reﬁ nement of their previous protocol,

whereby high-quality embryos could be obtained. Based on a two-

step culture system, they produced MDEs that germinated into

haploid or DH plants at a rate higher than 95 % [

230 ]. With respect

to sweet pepper types, in the last years several reports showed that

it is also possible to produce DHs through microspore culture,

although not as easy as it seems to be in hot types ([

191 , 219 , 231 ]

and our unpublished results). However, the problem of embryo

quality and ability to germinate still appears as a major bottleneck

to be overcome.

In summary, at present there are four main types of protocols

shown to experimentally induce microspore embryogenesis with

an acceptable, although variable efﬁ ciency: (1) the Dumas de Vaulx

method, (2) the biphasic method of Dolcet-Sanjuan, (3) the shed

microspore method of Supena, and (

4 ) the isolated microspore

method. The four have proven useful in obtaining DH pepper

plants. However, not all of them are in principle applicable to all

genotypes. In fact, the genotype of the donor plant is one of the

most decisive factors in the induction of pepper androgenesis [

180 ,

182 – 186 , 189 , 224 , 227 ]. In some cases, the optimization of

growth conditions for a given genotype is not sufﬁ cient to over-

come the barriers imposed by the genotype itself [

186 ]. However,

the possibility of applying different types of inductive protocols

allows for the choice of the most convenient for each variety. For

example, the cultivars “Quito” and “Piquillo” show a null/very

low response, respectively, to the method of Dumas de Vaulx [

189 ,

190 ], but “Quito” shows a fairly acceptable response to the shed-

microspore method [

190 ] and “Piquillo” shows a positive response

to the biphasic method of Dolcet-Sanjuan [

227 ]. Therefore,

before starting a breeding program based on DH production it is

advisable to assess the response of each variety to the different

types of induction protocol available. The development of the hap-

loid embryo is a second major drawback for an efﬁ cient DH produc-

tion in pepper. It seems that for most of the published methods, a

signiﬁ cant amount of MDEs are lost during the transition of a pro-

liferating, yet undifferentiated globular embryo into a heart-shaped

Jose M. Seguí-Simarro

233

bilateral one. Problems in the formation of the shoot apical meristem

and the cotyledons sometimes constrain a proper germination of

the embryo. It would be advisable to devote more efforts to the

knowledge of the particular developmental requirements of these

embryos, in order to facilitate their transition towards a mature

embryo.

8 Other Solanaceae

Aside of Datura and the ﬁ ve main solanaceous crops, attempts

have been made in other Solanaceae to obtain haploids and DHs

via anther or microspore culture . The following list illustrates some

representative examples: Atropa belladonna [ 232 – 234 ], Hyoscyamus

niger [

235 – 241 ], Hyoscyamus muticus [ 242 ], Petunia sp . [ 233 ,

243 – 251 ], Physalis ixocarpa [ 252 , 253 ], Solanum bulbocastanum

[

254 ], Solanum dulcamara [ 255 , 256 ], Solanum iopetalum [ 257 ],

Solanum surattense [

258 ], Solanum torvum [ 259 ], and Solanum

viarum [

260 ]. Although more studies will surely be published in

the future in these and other solanaceous species, the lower impor-

tance of them, compared to the ﬁ ve major solanaceous crops,

makes it unlikely to expect major achievements related to these

other Solanaceae.

9 Notes

1. Due to the signiﬁ cance of their ﬁ nding, both in fundamental

and applied terms, it is interesting to add some bits of history to

this review about the “making off” of this discovery. According

to a recent e-mail exchange that the author had with Dr.

Maheshwari, his discovery, as many others in experimental sci-

ences, came quite by an accident. The story starts with the

father of Satish Maheshwari, named Panchanan Maheshwari. In

his young days at Harvard in 1945, Panchanan Maheshwari

came into contact with A.F. Blakeslee, who inspired him to

investigate on haploid production. At that time it made no

sense to think of pollen as a source of haploid embryos, so all

efforts were directed to ovule culture. Indeed, Dr. Maheshwari’s

wife, Nirmala, was a former Ph.D. student at the lab of his

father (Panchanan Maheshwari), where they established a tech-

nique for ovule culture. In 1960, Satish Maheshwari and his

wife were working as postdoctoral researchers at the California

Institute of Technology (USA). In the James Bonner labora-

tory, they participated in the discovery of the existence of RNA

polymerases in plant nuclei and chloroplasts. Then, he returned

to India, to the Botany Department at Delhi University, headed

at that time by his father. Satish Maheshwari did not know

Androgenesis in Solanaceae

234

exactly what to do, since the continuation of his previous work

on nucleic acids was very difﬁ cult in a Department which by

that time had only microscopes, microtomes, and tissue culture

facilities. Following the advice of his former mentor (James

Bonner), he wanted to focus on fundamental research. In an

attempt to combine his research interests with the available

equipment, he decided to investigate the control of meiosis in

anther culture s. The idea was to culture anthers, then isolate

young microspore mother cells and see whether hormonal or

physical treatments were able to turn meiosis into mitosis or

vice versa. At this point Sipra Guha joined his group as a post-

doctoral researcher. They chose Datura innoxia for two very

simple reasons: by the time he started this project, there were

Datura plants ﬂ owering in the Botanical Garden of his univer-

sity, and due to the large size of the anthers, this seemed to be

the right material for his research. When they started the proj-

ect, accidentally, they discovered microspore embryogenesis ,

publishing their results in Nature in 1964. In words of Dr.

Maheshwari, “ we were naïve to try to study molecular biology

of meiosis this way, but then it led to the happy accident of

embryos popping out of anthers ” . After the discovery, it was

very difﬁ cult for Dr. Maheshwari to believe that the embryos

were originated from pollen grains, and that they were haploid.

Sipra Guha had to convince him providing the scientiﬁ c

evidences that led to the publication of their second Nature

paper in 1966 [

12 ].

Acknowledgments

Thanks are due to Dr. Satish C. Maheshwari for sharing with the

author some bits of the history of Datura anther culture s, and to

Mr. Edgar García for the acquisition of some of the pictures shown

in this review. This work was supported by grant AGL2014-55177

from Spanish MINECO to J.M.S.S.

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_10, © Springer Science+Business Media New York 2016

Chapter 10

Bioreactors for Plant Embryogenesis and Beyond

Liwen Fei and Pamela Weathers

Abstract

A variety of different bioreactors have been developed for use in initiating and cultivating somatic embryos.

The various designs for embryogenesis and culture are critically evaluated here. Bioreactor optimization and

operation methods are also described along with recommendations for use based on desired outcome.

Key words Balloon-type bubble reactor , Mist reactor , RITA

® , Temporary immersion system , TIS ,

Wave reactor

1 Bioreactor Types for Initiating and Cultivating Somatic Embryos

and Resulting Plantlets

There are many types of bioreactors that have been designed and

used for cultivating plant cell, tissue and organ cultures as illus-

trated schematically in Fig.

1 . These include: (a) classic stirred tank

(STR); (b) bubble column reactor ( BCR ); (c) balloon-type bubble

reactor ( BTBR ); (d) airlift reactor; (e) temporary immersion system

( TIS ) reactor; (f) RITA

® (a variation of TIS); (g) rotating drum; (h)

life reactor; (i) bag lined BCR; (j) bag lined STR; (k) wave reactor ;

(l) undertow reactor; (m) box-in-a-bag; and (n) mist reactor . A

number of these reactors also have disposable culture bags whereby

both contamination risk and initial capital cost are reduced. In Fig.

1 ,

reactors with disposable culture bags include: (h) life reactor; (i)

bag lined BCR; (j) bag lined STR; (k) wave reactor; (l) undertow

reactor; (m) box-in-a-bag; and (n) mist reactor. Bioreactors are

generally divided into two main groups, liquid phase and gas phase,

and have been used with varying success. For a more in depth

description of the utility of these different types of reactors for

in vitro cultivation of a variety of different plant species, see recent

reviews by Paek et al. [

1 ] and Weathers et al. [ 2 , 3 ]. Here we focus

on the key conditions of concern to embryo culture and character-

istics of these different types of reactors, as they have been used

speciﬁ cally for somatic embryo (SE) initiation and cultivation.

246

2 Conditions Critical to Formation and Culture of SEs in Reactors

All plant cell cultures require media selection and optimization per

species, and embryogenic cultures are no different. Bioreactors

offer some distinct advantages in the larger volume cultivation of

in vitro cells, tissues, or organs in that gases and nutrients can be

added or removed at various stages of cultivation. However, the

type of reactor will also dictate how such environmental factors are

controlled. For example, a liquid-based system like a BCR is usu-

ally run in a batch mode and the nutrient medium is not externally

altered. On the other hand, gas composition and delivery rate in a

BCR are critical and must be optimized to maximize SE generation

and development. Operating parameters, beyond nutrient medium

constituents, that are of considerable concern and that can be most

fully controlled in bioreactors are: gases and gas exchange, shear

stress , and light.

vent

vent vent vent

air

air air

air

valve

media

gas

Inoculum

sample

bag

rocking unit

Magnetic stirrer

gas/media exit

a

fg

bc d e

hij

k

mn

l

Fig. 1 Various bioreactors used for plant cell, tissue, organ and somatic cell embryogenesis

Liwen Fei and Pamela Weathers

247

There are three main gases of concern for any plant culture: O

2 ,

CO

2 , and ethylene (C

2 H

4 ). In liquid cultures, gases have limited

solubility, so efﬁ cient gas transfer is challenging. Gas solubility is a

function of temperature, pressure and solutes such as salts and sug-

ars [

4 , 5 ]. For the gases most important to plant cultures, solubility

decreases as temperature and solutes increase. At 25 °C the water

solubility of CO

2 and C

2 H

4 is about 25 and 4 times higher, respec-

tively, than O

2 , so in liquid phase reactors the amount of gas avail-

able to a growing tissue culture is quite restricted. Oxygen affects

differentiation of embryogenic cells. Although callus formation

and explant viability is not affected, anoxia almost completely

inhibits embryogenesis [

6 ], suggesting oxygen is required for

embryogenesis. Low O

2 concentration may enhance embryo for-

mation by simulating the in ovule environment normally encoun-

tered during zygotic embryo development [

7 ]. The overall demand

on O

2 then increases during subsequent maturation to the cotyle-

don stage [

8 – 10 ].

Elevating CO

2 level in the culture container often improves

somatic embryo genesis [

11 , 12 ] and effective concentration varies

from 0.3 to 5 % with species and cultivars [

11 – 16 ]. An extremely

high CO

2 concentration (10 %), however, is toxic to embryo prolif-

eration [

17 ]. Of course O

2 and CO

2 are not present, nor do they

affect, plant cells in isolation. They function in combinations that

may ﬂ uctuate at different optimal concentration, depending on the

developmental stage. Thus, while the early stage of embryogenesis

may prefer relatively low O

2 and high CO

2 levels [ 6 , 18 – 20 ], there

is considerable species and cultivar variation. For example, embryo

initiation of celery is favored under 30 % dissolved oxygen (DO,

about 5 mL/L) plus 3 % CO

2 [ 15 ]. On the other hand one cultivar

of Cyclamen persicum had signiﬁ cantly more embryos formed at 40

% DO (about 7 mL/L) than another cultivar where there was better

embryo formation at lower oxygen levels [

21 ]. Embryo differentia-

tion is also affected as shown in Coffea arabica where a DO of 80 %

(about 14 mL/L) generated more total embryos, but many fewer

at the torpedo stage than at 50 % DO (about 8.4 mL/L) [

19 ].

Ethylene also seems to be required for early differentiation

during somatic embryo genesis [

15 , 22 – 28 ]. However, there are

some conﬂ icting reports on the effect of headspace C

2 H

4 on

somatic embryogenesis [

29 , 30 ]. It is thus likely that species and

cultivars vary in their endogenous production of C

2 H

4 and opti-

mal C

2 H

4 concentration for embryo development . Sub-optimal-

producers may need an exogenous supply of C

2 H

4 , while

over- producers may require removal of C

2 H

4 [ 18 , 31 , 32 ]. In

sealed containers C

2 H

4 generally accumulates to toxic levels for

subsequent embryo maturation [

33 ]. Embryo development can

be improved, however, by increasing ventilation, using a C

2 H

4

trap (e.g., potassium permanganate), adding inhibitors of C

2 H

4

biosynthesis (e.g., aminooxyacetic acid, aminoethoxyvinylgly-

cine), or action (e.g., silver nitrate, CO

2 ) [ 34 – 39 ].

2.1 Gases

Bioreactors for Plant Embryogenesis

248

Although O

2 , CO

2 , and C

2 H

4 are the key metabolic gases of

concern for plant and SE culture, other gases may also be impor-

tant. For example, ozone (O

3 ) is a strong oxidant used to disinfect

water. O

3 readily decomposes to O

2 with no toxic by-product, so it

could conceivably be used in reactors as a periodic in situ disinfec-

tant to help maintain SE or other plant tissues in axenic culture.

Indeed, Aloe barbadensis grown in a 4 L BTBR with 1–15 min

daily treatment of O

3 over 4 weeks of culture responded quite

favorably [

40 ]. Although O

3 can be toxic to plants, limited inter-

mittent use may be a reasonable process option.

The gas environment inside a culture container, as well as in a bio-

reactor , is highly dependent on the gas exchange rate (ventilation)

of the headspace gas. Increasing gas exchange beneﬁ ts growth by

increasing CO

2 level and reducing relative humidity, as well as toxic

volatiles (i.e., C

2 H

4 ) in the headspace of a culture container [ 41 –

43 ]. In practice, the gas exchange rate can be increased by increas-

ing passive ventilation in culture containers or integrating a system

of forced ventilation. In passive ventilated culture containers, gas

exchange rate is increased by using porous closures or gas perme-

able membranes on the closure [

44 – 47 ]. By using these strategies,

the gas exchange rate can be elevated from 0.04 times/h (0.00066

vvm) under non-ventilated conditions to around 5 times/h (0.083

vvm) [

43 , 48 , 49 ]. Forced ventilation is best described as vvm,

which is deﬁ ned as the number of volumetric exchanges of head-

space gas per unit time (e.g., per minute) within the culture vessel.

Use of vvm allows for comparison between differently ventilated

culture systems. Except for small culture containers, the gas

exchange rate under passive ventilation may still be limited, so even

in spite of CO

2 enrichment, CO

2 concentration inside the con-

tainer is challenging to maintain at ambient levels (0.039 %) [

42 ].

Increasing ventilation in gelled medium is also limited by water

potential, as the medium can desiccate when gas exchange is

increased even via passive ventilation, and in vitro growth may

therefore become limited [

50 , 51 ]. Forced ventilation, on the

other hand, is more effective than passive ventilation in terms of

gas exchange rate for promoting photosynthesis of cotyledonary

stage embryos and their subsequent germination , as well as the

conversion to plantlets, in bioreactors [

52 – 54 ]. It is also a more

reliable means for controlling the gas environment inside culture

container than passive ventilation [

55 ]. Forced ventilation is essen-

tial for maintaining efﬁ cient gas exchange for photoautotrophic

growth in large culture containers, i.e., bioreactors [

56 ]. Forced

ventilation is achieved by ﬂ ushing humidiﬁ ed air via an air pump,

connected to a sterile air ﬁ lter, into the culture container [

57 ]. The

gas exchange rate under forced ventilation can be adjusted to more

than 10 times/h (0.16 vvm), which efﬁ ciently replenishes CO

2 for

photosynthesis in bioreactors [

42 , 58 – 60 ] and provides more uni-

2.2 Gas

Exchange Rate

Liwen Fei and Pamela Weathers

249

form gas distribution in the headspace and, therefore, in the culture

medium. In forced ventilation, the greater the ventilation rate, the

more the gaseous environment in vitro is similar to that of the ex

vitro environment and, as a consequence, the more closely in vitro

plantlets resemble ex vitro plants in their morphophysiological

traits [

60 ].

To obtain adequate mass transfer of gases into large volumes of

liquid, usually requires signiﬁ cant agitation of the culture medium,

and this can induce hydrodynamic shear stress on the growing cells

or tissues. SEs are subject to shear stress and, to reduce shear forces,

a slow-speed “string bioreactor ” with bubble-free aeration, deliv-

ered by thin silicone tubing hanging inside the periphery of the

reactor, has been developed. This modiﬁ ed stirred tank reactor

(Fig.

1a ) has been used to propagate somatic embryo s of carrot ,

Norway spruce , birch, cyclamen, as well as shoots of Christmas

begonia with minimum or no damage from shear [

61 ]. There are

other stirred reactors similar to the stirred tank (Fig

1a ), but they

use a spinning ﬁ lter or cell lift impeller and aeration tubes to pro-

vide low shear and bubble free aeration culture of somatic embryos

[

62 – 64 ]. Low-shear mixing and aeration in reactors can also be

achieved by bubbling and these bioreactors include bubble column

( BCR ; Fig.

1b ), balloon type bubble ( BTBR ; Fig. 1c ) and air-lift

(Fig.

1d ) reactors. These reactors use air sparged into the bottom

of the reactor to create rising bubbles for mixing and gas diffusion.

Compared to bubble reactor s, the air-lift reactor has an additional

draft tube. However, the major disadvantage of air-lift and bubble

column reactors is foaming induced by large volumes of air bub-

bling, growth of plant tissue in the head space and loss of culture

medium volume. To overcome the foaming problem, various reac-

tor designs with a larger top-section diameter have been devel-

oped, and the BTBR appears to yield the best biomass due to its

high oxygen transfer coefﬁ cient [

65 ]. Volume loss is minimized

through humidiﬁ cation of the incoming gas by use, for example, of

Naﬁ on tubing (Perma Pure, Toms River, NJ, USA).

Although seemingly sparsely studied, light quality can also play a

role in embryogenesis. For example, compared to darkness, far-

red, or red-far-red exposure, red light induced a fourfold increase

in SEs of quince ( Cydonia oblonga ) [

66 ]. In contrast, initiation of

SEs from Agave tequilana showed no dependency on light quality,

but later development into the cotyledon stage was maximized

after exposure to either red or white light [

67 ]. In carrot cell sus-

pension s, darkness produced the most SEs, and did not differ from

cells exposed to red or green light [

68 ]. On the other hand, both

white and blue light inhibited SE formation [

68 ]. Michler and

Lineberger also showed that, in carrot, red light enhanced heart

stage SEs [

68 ]. Although light quality may have beneﬁ cial effects

2.3 Shear Stress

2.4 Light Effects

Bioreactors for Plant Embryogenesis

250

on embryogenesis, once the cell density in a liquid-phase reactor

increases, light penetration into reactors becomes a nontrivial task,

so darkness would certainly be the preferred condition for cultiva-

tion [

68 ]. Indeed, the box-in-a-bag reactor (Fig. 1m ) was devel-

oped because of the challenge in getting adequate light into

cylindrical TIS reactors [

53 , 54 , 69 ].

3 Liquid Phase Bioreactors for Embryos

Most bioreactors used for SE formation are liquid phase wherein

the embryos are immersed in liquid medium all the time or inter-

mittently. Here we describe those that have been used successfully

for somatic embryo genesis and are also commercially available

(Table

1 ). Although shake ﬂ asks can be used as small-scale bioreac-

tors, the ability to control gas delivery and shear can enhance pro-

ductivity of SEs. These reactors range in size from 15 mL to

20,000 L. Similar to a stirred tank (Fig.

1a ), the miniPerm

® is a

small-volume mechanically driven modular reactor that offers mul-

tiple, tiny culture systems, each of which can be fed a stream of gas.

Modularity provides the ﬂ exibility of testing multiple cell lineages

under simultaneous conditions but with small volumes and inocu-

lum. The wave reactor (Fig.

1k ) is basically a horizontal, transpar-

ent plastic bag, residing on a slowly rocking platform that provides

agitation of the liquid medium. As a result of the large surface area

of the liquid in the bag, there is relatively good gas exchange

potential; gases are provided either passively or actively via sterile

permeable ﬁ lters or membranes. Unfortunately the wave reactor

scales laterally, requiring a considerable footprint, instead of less

costly vertical space. To our knowledge there are few, if any, reports

of either the miniPerm

® or the wave reactor being used for SE

production.

Table 1

Commercial bioreactors currently available for embryo cultivation

Reactor type Current manufacturer Max. vol (L)

Mechanically driven membrane

bioreactor— miniPerm

® Sartorius AG, GDR and USA 0.015

Wave Bioreactor GE Healthcare Life Sciences, USA 0.3–500

BIOSTAT (wave type) Sartorius AG 1–600

Balloon-type bubble reactor Samsung Science Co., Seoul, So. Korea 4–20,000

TIS ( RITA

® , Plantima, Plantform) Vitropic, France; Plantima A-tech; Toronado,

Netherlands; Plantform, Sweden 1–4

TIS temporary immersion system

Liwen Fei and Pamela Weathers

251

Although not commercially available, the bubble column reactor

( BCR ; Fig.

1b ) is easy to construct in-house. It is comprised of a

cylinder of glass or plastic (autoclavable) with a bottom frit attached

via tubing to a gas supply, e.g., air, which passes through the frit,

subsequently forming small bubbles that rise through the column

of liquid medium, thereby aerating and mixing the culture. Gas

vents via a sterile ﬁ lter at the top. Many different species of

SE-forming plants have been grown in BCRs, with some examples

reported in Table

2 . Unfortunately one of the problems with BCRs

is foaming, and because of it the balloon-type bubble reactor

( BTBR ) was developed (Fig.

1c ). The broad surface area of the

culture liquid alleviates foaming and provides even better gas

exchange than the BCR. A variety of different plant species have

produced SEs in the BTBR at a variety of volumes (Table

2 ; Fig. 2 ).

Although the BTBR has been scaled to 500 L, it is constructed of

glass and at large scale must be provided with a stainless steel

superstructure, which adds to capital costs. Smaller scale versions

(e.g., 4 L) of the BTBR are, however, quite competitive with other

smaller bioreactors.

The TIS (Fig.

1e ) allows for periodic wetting of the inoculum

with nutrient medium. Liquid is pressure fed via tubing from the

bottom chamber into the top growth chamber to the level of the

inoculum that is located on a porous platform. Gas vents with ster-

ile ﬁ lters are used to equalize pressure. The liquid is held in the top

chamber for a short period of time and then drained back to the

bottom chamber until the next ﬁ lling. This can occur at any regu-

larly set interval, which is often species-speciﬁ c. TIS can easily be

built in-house using Nalgene bottles or other small vessels. A simi-

lar version is the “twin ﬂ asks” system (not shown) where liquid is

passed between two ﬂ asks horizontally, instead of vertically; this

system is also easy to build in-house. A commercially available

example of a TIS is the RITA

® system (about 1 L of total volume).

To initiate an immersion cycle, pressure is applied to the lower

chamber, pushing the medium into the upper chamber. This way

plant material is immersed in the bubbling medium, so providing

gentle mixing and headspace gas renewal. When the pressure is

released, the medium drains back to the lower chamber to com-

plete the immersion cycle. There is a diffusive aeration outlet on

the top of the apparatus to balance pressure.

As SEs mature and become chlorophyllous in the cotyledon-

ary stage, light transmittance becomes important for germination

of SEs and their subsequent plantlet development [

70 ]. However,

the cylindrically shaped vessels in most TIS restrict light penetra-

tion into their center, thus also restricting SE development in the

center of the culture vessel [

54 ]. To obtain uniform light transmit-

tance and, therefore, more or less synchronized SE maturation,

the box-in- a-bag TIS was developed for production of pre-germi-

nated embryos using torpedo stage embryos as inoculum [

53 , 71 ].

Bioreactors for Plant Embryogenesis

252

The growth chamber of this bioreactor is made of a transparent

disposable plastic bag, ﬁ tted outside a box with a lateral screen on

which SEs reside and grow. Despite the high light transmittance,

this bioreactor has a large footprint; it also has a problem in medium

mixing and the sterile vent connector parts on the bag are too costly

to be disposable [

69 ]. RITA

® and other types of TIS have been

used to culture SEs of many plant species, as noted in Table

2 .

4 Gas Phase Bioreactors for Embryos

The mist reactor is, to our knowledge, the only gas-phase reactor.

In this system, nutrient medium is provided to cells, explants tis-

sues or organs via an ultrasonic nozzle that yields a ﬁ ne mist,

coalescing and dripping back into the medium reservoir. Although

not commercially available, the mist reactor can readily be con-

structed in-house and used for experiments [

72 ]. It has a dispos-

able transparent plastic bag that is used as a culture chamber.

Recently, the mist reactor was used to generate carrot SEs [

73 ].

Cells were manually inoculated onto poly- l -lysine (PLL)-coated

nylon mesh that was then hung inside the reactor bag and grown

in misted medium for several weeks. Carrot cells were attached to

the PLL strips and formed SEs that developed into small rooted

Table 2

Some examples of SEs successfully cultivated in bioreactors.

Bioreactor type Species Volume (L) Ref.

BTBR Eleutherococcus senticosus 500 [ 65 ]

Transgenic E. senticosus SEs 130 [

77 ]

Panax notoginseng 3 [

78 ]

Musa acuminata cv Berangan (AAA) 5 [

79 ]

Santalum album 10 [

80 ]

TIS Coffea arabica 1

1–10 [

76 ]

[

54 ]

Saccharum spp. cv Q165 ≤1 [

81 ]

Hevea brasiliensis 1 [

82 ]

Theobroma cacao 1 [

83 ]

Bactris gasipaes Kunth 1 [

84 ]

BCR Castenea dentate x mollisima 0.1–1.0 [

85 ]

Eleutherococcus senticosus 10 [

86 ]

Lilium x formolongi (5 cvs) 2 [

87 ]

Picea sitchensis 2 [

88 ]

Mist Daucus carota 4 [ 73 ]

Shake ﬂ asks Quercus suber L. 0.1–0.25 [ 75 ]

BCR bubble column reactor, BTBR balloon type bubble reactor , SE somatic embryo s, TIS temporary immersion

system

Liwen Fei and Pamela Weathers

253

plantlets, while still hanging on the strips inside the reactor.

Subsequently, inoculation was attempted by spraying the cells

through the ultrasonic nozzle such that they landed on the hang-

ing PLL-coated nylon mesh. Cells attached, remaining for several

weeks while they developed into rooted plantlets. This method

suggested that fully developed plantlets, ready for planting into

soil, can be obtained in a single step in a bioreactor from cells that

underwent somatic embryo genesis.

Fig. 2 Large scale cultivation of Siberian ginseng in a balloon type bubble column reactor. Suspension cultures

of somatic embryo s of Eleutherococcus senticosus . ( a ) Embryogenic cells in MS liquid medium supplemented

with 30 g/L sucrose and 1 mg/L 2,4-dichlorophenoxyacetic acid. ( b ) Embryogenic suspension in 3-L capacity

balloon- type airlift bioreactor containing 2-L MS medium with 30 g/L sucrose. ( c ) Embryogenic suspension in

500 L balloon-type airlift bioreactor. ( d ) Biomass harvested from 500 L balloon bioreactor after 30 days of

culture (reproduced from [ 65 ] with permission from Springer Science + Business Media)

Bioreactors for Plant Embryogenesis

254

5 Scaling Up Somatic Embryogenesis and Somatic Embryo Cultivation

Before scaling up, all conditions, especially soluble components

like nutrients and plant growth regulators, should ﬁ rst be opti-

mized in small scale shake ﬂ asks. The move to larger scale reactors

should be made with the following important caveat: not all

responses can be directly scaled from shake ﬂ asks to any reactor

system, mainly because ﬂ asks and reactors usually differ signiﬁ -

cantly in design and operation. There are several reports that docu-

ment simple, yet effective, approaches for optimizing inoculum,

DO, and shear for scaling up SEs. For example, shake ﬂ asks were

used to optimize inoculum for SE formation in Coffea sp. [

54 ].

Inoculum densities of 0.1–3.0 g FW/L of suspension cells were

grown in 25 mL of medium in 250 mL Erlenmeyer ﬂ asks. If the

medium was refreshed with new medium each week, maximum

SEs were obtained at the 1.5 g inoculum level, with 25 % reaching

the torpedo stage; when medium was not refreshed the number of

SEs was much less. Although fewer SEs were obtained at 1.0 and

0.5 g FW/L inoculum, conversion rate to the torpedo stage was

considerably greater than at the higher inoculum density. At 3.0 g

FW/L, only a few SEs appeared, suggesting that high inoculum

density was inhibitory. In another study using cell suspension inoc-

ulum in 25 mL of medium in 250 mL Erlenmeyer ﬂ asks, banana

(hybrid FHIA-18 AAAB) also had an optimum inoculation den-

sity. Both inoculum density and culture medium were optimized

for somatic embryo production [

74 ].

In another instance, in Quercus suber three different shake

ﬂ asks were used to determine the optimum DO and shear condi-

tions for SE formation [

75 ]. The three types of shake ﬂ asks were: a

100 mL Erlenmeyer (EF100), a bafﬂ ed 150 mL Erlenmeyer

(BEF150), and a 250 mL Erlenmeyer (EF250). In conjunction

with three orbiting speeds of 60, 110, and 160 rpm, they were able

to vary the oxygen volumetric mass transfer coefﬁ cient ( K

L a /h)

more than tenfold, from 0.11 in the EF100 at 60 rpm to 1.47 in

the BEF150 at 160 rpm with an oxygen transfer rate that increased

sixfold at the higher orbit speed. Similarly the shear force index

(SFI, cm/min) was altered from 1.4 × 10

3 in the EF100 at 60 rpm

to 8.8 × 10 3 in the BEF150 at 160 rpm. These simple shake ﬂ ask

studies provided statistically signiﬁ cant differences that enabled

relatively easy optimization of Q. suber for embryogenesis, with the

largest number of embryogenic cell clumps obtained in the BEF150

at 160 rpm.

In contrast to the production of plantlets, hairy roots or pro-

duction of secondary metabolites, scaling up embryogenesis for

cells has the end goal of working with smaller volume reactors. A

considerable number of viable embryos can be produced within a

relatively small reactor volume, even in a shake ﬂ ask. For example,

1,600 Coffea embryos were obtained in the 1 L volume of one

Liwen Fei and Pamela Weathers

255

RITA

® container [ 76 ]. However, for large plantations of elite

genotypes, larger volumes may be required, and thus far the

balloon- type bubble reactor has shown considerable promise.

Recently, in South Korea, Shohael et al. [

65 ] scaled up produc-

tion of Siberian ginseng SEs ( Eleutherococcus senticosus ) to 500 L in

a balloon-type reactor (Fig.

2 ). They inoculated somatic embryo s

grown ﬁ rst in shake ﬂ asks into a 3 L balloon feeder reactor that

then was fed into the ﬁ nal 500 L reactor. Using four different 3 L

scale (2 L working volume) reactor designs, the balloon reactor

that performed best, in terms of biomass and eleutheroside yield,

was determined, showing that a stepped aeration protocol of 0.05–

0.3 vvm and 5 g cells/L of inoculum density was optimal. Finally,

they compared large scale production in 500 L rotating drum and

balloon reactors, and the balloon outperformed the rotating drum.

6 Conclusions

There are a number of different reactor types that can be used

for the cultivation of embryogenic cell lines. Some involve use

of a disposable culture bag, while several are simple and easy to

construct in-house. However, only a few are commercially avail-

able. Thus, although large scale production of somatic embryo s in

bioreactors is certainly possible, the possibility of commercial

exploitation will clearly be the driving force in the further develop-

ment of the technology. Furthermore, each plant species needs to

be separately optimized for reactor design and production operat-

ing parameters.

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Bioreactors for Plant Embryogenesis

Part II

Protocols of Somatic Embryogenesis in Selected

Important Horticultural Plants

263

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_11, © Springer Science+Business Media New York 2016

Chapter 11

Somatic Embryogenesis and Genetic Modiﬁ cation of Vitis

Sadanand A. Dhekney , Zhijian T. Li , Trudi N. L. Grant , and Dennis J. Gray

Abstract

Grapevine embryogenic cultures are ideal target tissues for inserting desired traits of interest and improving

existing cultivars via precision breeding (PB). PB is a new approach that, like conventional breeding, utilizes

only DNA fragments obtained from sexually compatible grapevine plants. Embryogenic culture induction

occurs by placing leaves or stamens and pistils on induction medium with a dark/light photoperiod cycle

for 12–16 weeks. Resulting cultures produce sectors of embryogenic and non-embryogenic callus, which

can be identiﬁ ed on the basis of callus morphology and color. Somatic embryo development occurs follow-

ing transfer of embryogenic callus to development medium and cultures can be maintained for extended

periods of time by transfer of the proliferating proembryonic masses to fresh medium at 4–6- week intervals.

To demonstrate plant recovery via PB, somatic embryos at the mid-cotyledonary stage are cocultivated with

Agrobacterium containing the desired gene of interest along with a, non-PB, enhanced green ﬂ uorescent

protein/neomycin phosphotransferase II (e gfp/nptII ) fusion gene. Modiﬁ ed cultures are grown on prolif-

eration and development medium to produce uniformly modiﬁ ed somatic embryos via secondary embryo-

genesis. Modiﬁ ed embryos identiﬁ ed on the basis of green ﬂ uorescence and kanamycin resistance are

transferred to germination medium for plant development. The resulting plants are considered to prototype

examples of the PB approach, since they contain egfp/nptII , a non-grapevine-derived fusion gene. Uniform

green ﬂ uorescent protein (GFP) ﬂ uorescence can be observed in all tissues of regenerated plants.

Key words Agrobacterium , Culture medium , Embryogenic cultures , Growth regulators , Plant tissue

culture , Precision breeding , Vitis

1 Introduction

Grapevine is highly prized and grown worldwide for consumption

as fresh fruit and processed products, including jam, jelly, juice, raisin,

and, particularly, wine. Grape and its products contain a number of

ﬂ avonoid and non-ﬂ avonoid phenols that act as antioxidants and

impart health-beneﬁ cial properties. Resveratrol and proanthocy-

anidins present at high levels in wine possess anti-inﬂ ammatory

activities and are responsible for cardioprotection [

1 ]. A number of

grapevine cultivars have been cultivated for centuries and are greatly

valued for their speciﬁ c fruit/enological characteristics. Only 35

elite, mostly ancient, grapevine cultivars account for 66 % of acreage

264

worldwide, as consumers continually seek out the wines produced

from them [

2 ]. However, the elite cultivars, having been selected in

antiquity with no directed genetic improvement possible since, are

susceptible to a number of fungal, bacterial, and viral diseases; they

require substantial chemical control in traditional production areas

and cannot be grown at all in regions with extreme climatic condi-

tions. Improving abiotic and biotic stress tolerance of these elite

cultivars by conventional breeding is impossible because their key

sensory attributes are invariably lost. For example, a prized strain of

“Pinot Noir” cannot be improved by conventional breeding to cre-

ate a disease resistant “Pinot Noir.” This is because grapevine, like

many woody- perennial crops, are out-crossers, exhibiting self-

incompatibility and inbreeding depression. New conventionally

bred varieties, despite being resistant and even producing accept-

able wine, never correspond to their elite counterparts, since exist-

ing enological characteristics are disrupted, thereby meeting

consumer recalcitrance [

3 – 5 ]. Inserting desired traits via precision

breeding technology is a viable alternative for improving elite

grapevine cultivars without altering their highly prized enological

characteristics [

6 , 7 ].

Recent advances in grapevine genome sequencing have fos-

tered discovery of desirable traits to instill into elite cultivars, while

still maintaining their unique varietal characteristics. Grapevine

improvement via precision breeding (PB) involves using DNA

sequences found solely in the grapevine genome and it is a logical

reﬁ nement of conventional breeding, only recently made possible

by advances in cell culture, gene insertion, and computational

technology [

7 – 9 ]. Grapevine embryogenic cultures have long been

the targets of choice for inserting genes encoding desired traits,

since single cells on their surface can be prompted to develop into

complete plants. Hence gene insertion into such totipotent cells

results in plants that stably express the desired trait [

10 , 11 ]. An

embryogenic response in a grapevine cultivar involves a complex

interaction of the genotype with explant, culture medium and cul-

ture conditions [

4 , 12 ]. This necessitates protocol optimization for

each grapevine cultivar grown in speciﬁ c regions of the world. We

have studied the embryogenic response of leaf and ﬂ oral explants

at various developmental stages, various media compositions, and

culture conditions for a large number of cultivars over the last three

decades [

13 , 14 ]. Embryogenic cultures are maintained on devel-

opment medium for extended periods of time by careful selection

and transfer of proliferating embryonic masses. Following somatic

embryo development and germination , regenerated plants are

hardened in a growth room and transferred to a greenhouse.

We also continue to optimize our previous protocols for gene

insertion by improving culture development, cocultivation proce-

dures, reducing culture necrosis, and increasing plant recovery.

Sadanand A. Dhekney et al.

265

Genetically modiﬁ ed plants have been recovered from a wide array

of Vitis species, cultivars, and interspeciﬁ c hybrids [

15 – 19 ].

This chapter describes speciﬁ c methods for the induction,

maintenance, and genetic modiﬁ cation of grapevine embryogenic

cultures to insert desired traits of interest in order to produce

precision- bred versions of elite cultivars (Fig.

1 ). Cultures are initi-

ated from leaves and/or ﬂ oral explants on a wide array of culture

media. Rapid proliferation of embryogenic cultures is obtained by

growing them in liquid medium [

20 ] and long term maintenance

is achieved by culture on specialized X6 medium [

12 ]. Somatic

embryo s at the mid-cotyledonary stage of development are cocul-

tivated with disarmed (non-disease causing) Agrobacterium har-

boring the desired genes of interest. For this demonstration,

modiﬁ ed cultures are identiﬁ ed on the basis of non-precision-bred

green ﬂ uorescence and kanamycin resistance. Plants obtained fol-

lowing germination of embryos are hardened in a growth room

and transferred to a greenhouse. The genetically modiﬁ ed status of

regenerated plants is conﬁ rmed by the uniform expression of the

green ﬂ uorescence protein gene in various plant tissues.

Fig. 1 Somatic embryo genic culture and genetic modiﬁ cation system for grapevine (reproduced from Ref. 26

with permission from Nature Publishing Group)

Somatic Embryogenesis in Vitis

266

2 Materials

1. Autoclave.

2. Bead sterilizer.

3. Stereomicroscope.

4. Fiber-optic illuminator.

5. Forceps.

6. Scalpels.

7. Clorox

® bleach or equivalent.

8. Sterile Whatman 3MM ﬁ lter paper.

9. Tween 20.

10. Sterile distilled water.

11. 100 × 15 mm plastic Petri dishes.

12. GA7 Magenta culture vessels.

13. Laminar airﬂ ow sterile culture hood.

14. Growth chamber.

15. pH meter.

16. Micropipettes and micropipette tips.

17. Spray bottles.

18. 125 mL Erlenmeyer ﬂ asks.

19. 960 μM sieves.

20. Rotary shaker.

21. Leica MZFLIII stereomicroscope or equivalent equipped for

epi-ﬂ uorescence with an HBO 100 W Mercury lamp illumina-

tor and a green ﬂ uorescent protein ( GFP ) ﬁ lter set composed

of an excitation ﬁ lter (470/40 nm), a dichromatic beam split-

ter (485 nm), and a barrier ﬁ lter (525/50 nm) (Leica

Microscopy System Ltd., Heerbrugg, Switzerland).

1. One-year-old, dormant grapevine cuttings ( see Note 1 ).

2. Established micropropagation cultures.

1. Embryogenic culture induction from leaf explants (NB2

medium): Nitsch and Nitsch [

21 ] macro-, micronutrients and

vitamins, 0.1 g/L myoinositol , 20 g/L sucrose , 1.0 μM ben-

zyl amino purine (BAP), 5.0 μM 2,4-dichlorophenoxyacetic

acid (2,4-D), 7.0 g/L Tissue culture (TC) grade Agar

(Phytotechnology labs), pH 6.0 ( see Note 2 ).

2. Embryogenic culture induction from stamen and pistil

explants (MSI medium): Murashige and Skoog [

22 ], macro-,

micro- nutrients and vitamins, 0.1 g/L myo-inositol , 20 g/L

2.1 Supplies

and Equipment

2.2 Explant Sources

2.3 Culture Medium

Composition

Sadanand A. Dhekney et al.

267

sucrose , 4.5 μM BAP, 5.0 μM 2,4-D, 7 g/L TC agar, pH 6.0

( see Note 3 ) [

13 ].

3. The following media may variously be used for embryogenic

culture induction from anther and pistil explants:

(a) PIV medium: Nitsch and Nitsch macro- and micronutri-

ents, B5 vitamins, 60 g/L sucrose , 8.9 μM BAP, 4.5 μM

2,4- D, 3.0 g/L Phytagel , pH 5.7 [

23 ].

(b) X1 medium: Modiﬁ ed MS macro-, micronutrients and

vitamins, which lack glycine and consisting of modiﬁ ed

MS nitrate (X nitrate) consisting of 3.033 g/L KNO

3 and

0.364 g/L NH

4 Cl, 0.1 g/L myoinositol , 20 g/L sucrose ,

5.0 μM BAP, 2.5 μM 2,4-D and 2.5 μM beta-naphthoxy-

acetic acid ( NOA ), 7 g/L TC agar, pH 5.8 [

13 ].

(c) X2 medium: Modiﬁ ed MS macro-, micronutrients and

vitamins, which lacks glycine and MS nitrate being

replaced with X nitrate consisting of 3.033 g/L KNO

3

and 0.364 g/L NH

4 Cl, 0.1 g/L myoinositol , 20 g/L

sucrose , 5.0 μM BAP, 15.0 μM 2,4-D and 2.5 μM NOA ,

7 g/L TC agar, pH 5.8 [

13 ].

(d) NI medium: Nitsch and Nitsch macro-, micronutrients

and vitamins, 0.1 g/L myoinositol , 20 g/L sucrose ,

5.0 μM BAP, 2.5 μM 2,4-D and 2.5 μM NOA , 7 g/L TC

agar, pH 5.8 [

13 ].

(e) NII medium: Nitsch and Nitsch macro-, micronutrients

and vitamins, 0.1 g/L myoinositol , 20 g/L sucrose ,

5.0 μM BAP, 15.0 μM 2,4-D and 2.5 μM NOA , 7 g/L

TC agar, pH 5.8 [

13 ].

4. Embryogenic culture maintenance in liquid medium (B5/ MS

medium ): B5 macro-nutrients, MS micronutrients and vita-

mins, 0.4 g/L glutamine , 60 g/L sucrose , 4.5 μM 2,4-D,

pH 5.8 [

19 ].

5. Embryo development and maintenance medium (X6 medium):

Modiﬁ ed MS macro-, micronutrients and vitamins, which

lacks glycine and MS nitrate being replaced with X nitrate con-

sisting of 3.033 g/L KNO

3 and 0.364 g/L NH

4 Cl, 60.0 g/L

sucrose , 1.0 g/L myoinositol , 7.0 g/L TC agar , 0.5 g/L

activated charcoal , pH 5.8 ( see Note 4 ).

6. Agrobacterium solid culture medium (YEP medium): 10 g/L

yeast extract, 10 g/L peptone, 5.0 g/L NaCl, 20 g/L agar ,

pH 7.0.

7. Agrobacterium liquid culture medium (MG/L medium):

5.0 g/L mannitol , 1.0 g/L

L -Glutamate, 5.0 g/L tryptone ,

2.5 g/L yeast extract, 5.0 g/L NaCl, 150.0 mg/L KH

2 PO

4 ,

100.0 mg/L MgSO

4 ·7H

2 O, 2.5 mL/L Fe–EDTA, pH 7.0

( see Note 5 ).

Somatic Embryogenesis in Vitis

268

8. Agrobacterium liquid transfer medium (X2 medium): X6

medium modiﬁ ed to contain 20.0 g/L sucrose without TC

agar and activated charcoal , pH 5.8.

9. Liquid cocultivation medium (DM medium): DKW basal salts

[

24 ], 0.3 g/L KNO

3 , 1.0 g/L myo -inositol , 2.0 mg/L each of

thiamine –HCl and glycine , 1.0 mg/L nicotinic acid , 30 g/L

sucrose , 5.0 μM BAP, 2.5 μM each NOA and 2,4-D, pH 5.7.

10. Callus induction medium (DM medium): DKW basal salts,

0.3 g/L KNO

3 , 1.0 g/L myo -inositol , 2.0 mg/L each of thia-

mine –HCl and glycine , 1.0 mg/L nicotinic acid , 30 g/L

sucrose , 5.0 μM BAP, 2.5 μM each NOA and 2,4-D, 7.0 g/L

TC agar , 200 mg/L each of carbenicillin and cefotaxime , and

100 mg/L kanamycin, pH 5.7.

11. Embryo germination medium (MS1B): MS macro-, micronu-

trients and vitamins, 0.1 g/L myoinositol , 30.0 g/L sucrose ,

1.0 μM BAP, 7.0 g/L TC agar , pH 5.8.

1. Rifampicin : Filter-sterilized stock solutions containing rifam-

picin at 20 mg/mL ( see Note 6 ).

2. Kanamycin sulfate: Filter-sterilized stock solutions containing

kanamycin sulfate at 100 mg/mL.

3. Carbenicillin and cefotaxime : Filter-sterilized stock solutions

containing either carbenicillin or cefotaxime at 200 mg/mL.

1. Binary vector containing the gene of interest and an egfp/

nptII fusion gene (reporter marker fusion) under the control

of a constitutive promoter.

2. Agrobacterium stock (containing the binary vector) stored in

glycerol at −70 °C.

3 Methods

Carry out all surface sterilization, explant isolation, and transfer

procedures using established aseptic techniques in a laminar airﬂ ow

hood. Clorox

® . Wrap all dishes with Paraﬁ lm

® .

1. Initiate in vitro micropropagation cultures from ﬁ eld- or

greenhouse- grown grapevine shoot tips ( see Note 7 ).

2. Excise unopened leaves, 1.5–5.0 mm in size, from in vitro-

grown micropropagation cultures and transfer them to Petri

dishes containing NB2 medium ( see Note 8 ).

3. Incubate cultures in darkness at 26 °C for 5–7 weeks for the

induction of embryogenic callus.

2.4 Antibiotic Stock

Solutions

2.5 Agrobacterium

Culture

3.1 Embryogenic

Culture Induction

from Leaf Explants

Sadanand A. Dhekney et al.

269

4. After 5–7 weeks, transfer cultures to light (65 μM m

−2 s

−1 and

16 h photoperiod) at 26 °C for 5 weeks. Screen callus cultures

for growth and possible contamination at weekly intervals.

5. Explants will produce callus cultures, which can be distin-

guished into cream-colored embryogenic callus and dark

brown non-embryogenic callus.

6. Carefully transfer the cream colored embryogenic callus to X6

medium for proliferation of proembryonic masses ( PEM ) and

development of somatic embryo s (SE).

1. Obtain grapevine inﬂ orescences from ﬁ eld-grown grapevines

or one year old dormant cuttings.

2. Surface-sterilize dormant cuttings in 25 % Clorox

® solution

with constant agitation for 5 min, followed by two washes

with sterile distilled water.

3. Make fresh cuts at the top and base of the cuttings and transfer

30 cm long cuttings to 500 mL conical ﬂ asks containing

250 mL sterile distilled water.

4. Transfer ﬂ asks under light (65 μM m

−2 s

−1 and 16-h photope-

riod) at 26 °C for 3–5 weeks for inﬂ orescence growth and

development.

5. Determine development stages of stamens and pistils using a

stereomicroscope to identify the optimum stage for the spe-

ciﬁ c cultivar ( see Note 9 ) (Fig.

2 ).

6. Surface-sterilize inﬂ orescences by rinsing brieﬂ y in 70 % etha-

nol followed by washing them in 25 % Clorox

® solution con-

taining a small drop Triton X-100 for 5 min with a periodic

manual high degree of agitation. Following washing with

Clorox

® solution, treat explants with three 5-min washes in

sterile distilled water.

7. Using a stereomicroscope, carefully excise intact stamens by

separating them from the calyptra and pistil. Place stamens

from ﬁ ve inﬂ orescences as a clump in the center of the Petri

dish containing induction medium and corresponding pistils

with the ﬁ lament stubs at the perimeter. Seal Petri dishes and

place in the dark at 26 °C for 5 weeks ( see Note 10 ).

8. After 5 weeks of incubation in the dark, transfer Petri dishes to

light (65 μM m

−2 s

−1 and 16-h photoperiod) at 26 °C. Screen

developing cultures using a dissecting microscope for the pres-

ence of embryogenic callus at weekly intervals for 12–16 weeks

( see Note 11 ).

9. Induction of embryogenic callus is observed either from the ﬁ la-

ment tip, connective tissue or in some cases from pistil explants.

10. Transfer the embryogenic callus to X6 medium for SE devel-

opment and proliferation.

3.2 Embryogenic

Culture Induction

from Stamen and Pistil

Explants

Somatic Embryogenesis in Vitis

270

1. Transfer 1.0 g rapidly growing embryogenic culture to sterile

125 mL Erlenmeyer ﬂ asks containing 40 mL autoclaved liquid

medium. Cover the ﬂ asks with aluminum foil and seal the neck

with Paraﬁ lm. Transfer the ﬂ asks to a rotary shaker and incu-

bate in diffused light (15 μM m

−2 s

−1 and 16-h photoperiod) at

120 rpm.

2. After 2 weeks, separate differentiated somatic embryo s by ﬁ l-

tering cultures through a sterile 960 μM stainless steel and

collecting the ﬁ ne fraction. Transfer the ﬁ ne fraction to fresh

liquid medium and differentiated SE to X6 medium for

embryo development .

3. Maintain suspension cultures by transfer to fresh liquid

medium at 2–3-week intervals ( see Note 12 ).

1. Transfer embryogenic cultures obtained from induction

medium to X6 medium for development and proliferation of

SE (Fig.

3a ).

3.3 Embryogenic

Culture Proliferation

in Liquid Medium

3.4 Embryogenic

Culture Maintenance

Fig. 2 Stamen and pistil explant developmental stages in Vitis . Four stages, I ( a , b ), II ( c , d ), III ( e , f ), and IV ( g ,

h ) can be identiﬁ ed on the basis of inﬂ orescence size, stamen color and size, and microspore development

stage. Stage I ﬂ ower clusters are about 2.5–3.0 cm long, individual ﬂ ower buds 0.5–0.7 mm in diameter,

anthers 0.1–0.2 mm in length, white in color and clear in appearance. Stage II ﬂ ower clusters are about

6–8 cm long with individual ﬂ ower buds being approximately 1.5 mm in diameter. Anthers are 0.8–1.0 mm

long, yellowish in color, and appear translucent with clear walls. Stage III ﬂ ower clusters are 9–10 cm long and

individual ﬂ ower buds 1.5–2.0 mm in diameter. Anthers are 1.0 mm in length, yellowish in color, and cloudy in

appearance with clear walls. The locule appears cloudy and yellowish in color. Microspore walls are thicker

and well developed. Stage IV ﬂ ower clusters are greater than 10 cm in length and individual ﬂ ower diameter

similar to Stage III. Anthers are 1.0 mm long and yellowish in color with completely opaque walls. The locule

appears yellow in color and opaque. Microspore walls are thicker and pores in the cell wall are evident (repro-

duced from Ref. 13 with permission from American Society of Horticultural Sciences)

Sadanand A. Dhekney et al.

271

2. Maintain embryogenic cultures by precisely separating PEM

from differentiated SE using a stereomicroscope as described

below (Fig.

3b, c ) and selectively transfer only PEM to fresh

X6 medium at 4–6-week intervals ( see Note 13 ) (Fig

3d, e ).

1. Streak Agrobacterium culture stock containing the binary

plasmid onto a Petri dish containing solid YEP medium with

20 mg/L rifampicin and 100 mg/L kanamycin. Incubate

dishes in the dark at 26 °C for 2–3 days until single bacterial

colonies are visible.

2. Transfer a single bacterial colony to a 125 mL conical ﬂ ask

containing 30 mL MG/L medium with 20 mg/L rifampicin

and 100 mg/L kanamycin. Seal the ﬂ ask with Paraﬁ lm and

incubate on a rotary shaker at 180 rpm and room temperature

for 16–20 h.

3.5 Agrobacterium

Culture Initiation

for Plant Genetic

Modiﬁ cation

Fig. 3 Embryogenic culture maintenance in Vitis . Actively proliferating proembryonic masses contained within

embryogenic culture masses grown on X6 medium ( a ) are identiﬁ ed, sub-cultured, and manipulated using a

stereomicroscope placed in a laminar ﬂ ow culture hood and illuminated with a ﬁ ber optic light source ( b , c ).

Only microscopic proembryonic tissue masses are selected ( d ) and these are accumulated so as to create ﬁ ve

cultures in each Petri dish containing 30 mL ( thick ) of freshly made X6 medium ( e ) and the cycle is repeated

at 4–6-week intervals. Proembryonic masses are uniformly composed of small, densely cytoplasmic embryo-

genic cells ( f ). It is important to keep a uniform subculture time and a stable incubation temperature to avoid

precocious germination

Somatic Embryogenesis in Vitis

272

3. Transfer the overnight culture to a 50 mL centrifuge tube and

spin at 4200 × g for 8 min at room temperature. Discard the

supernatant and resuspend the pellet in 20 mL liquid X2

medium. Transfer the culture to a 125 mL conical ﬂ ask and

incubate for an additional 3 h under the same conditions as

above. Use this culture for cocultivation .

1. Carefully transfer cotyledonary-stage SE to sterile Petri dishes.

Avoid wounding of embryos during transfer to prevent

culture browning ( see Note 14 ).

2. Add 5.0 mL Agrobacterium culture to the SE and mix thor-

oughly by swirling. Incubate for 7–10 min and then remove

the bacterial solution completely using a micropipette.

3. Transfer SE to a Petri dish containing two layers of ﬁ lter paper

soaked in liquid DM medium. Seal the Petri dish with Paraﬁ lm

®

and cocultivate in darkness at 26 °C for 72 h ( see Note 15 ).

4. Following cocultivation for 72 h, observe SE for transient

GFP expression using a stereomicroscope equipped for

epi-ﬂ uorescence.

5. Transfer cocultivated cultures to a 125 mL conical ﬂ ask con-

taining liquid DM medium with 200 mg/L each of carbenicil-

lin and cefotaxime , and 15 mg/L kanamycin.

6. Transfer the ﬂ ask to a rotary shaker at 110 rpm and wash SE

for 3 days to inhibit remnant bacterial growth.

7. Transfer washed cultures to each 100 × 15 mm Petri dish con-

taining 25 mL solid DM medium with 200 mg/L each of car-

benicillin and cefotaxime and 100 mg/L kanamycin.

8. Place Petri dishes in dark at 26 °C for 4 weeks to permit callus

development and proliferation.

9. After 4 weeks, transfer callus cultures to 100 × 15 mm Petri

dishes containing 30 mL X6 medium with 200 mg/L each of

carbenicillin and cefotaxime and 70 mg/L kanamycin for

secondary embryo development . Place Petri dishes in dark

and screen at weekly intervals for the presence of modiﬁ ed

SE lines.

10. Independent SE lines are identiﬁ ed by bright GFP ﬂ uores-

cence and kanamycin resistance ( see Note 16 ).

11. Transfer independent genetically modiﬁ ed embryo lines

to individual Petri dishes containing X6 medium with

200 mg/L each of carbenicillin and cefotaxime

and 70 mg/L

kanamycin.

12. Screen cultures for the development of modiﬁ ed embryo

development and proliferation.

3.6 Gene Insertion

into Embryogenic Cells

Sadanand A. Dhekney et al.

273

1. Transfer cotyledonary-stage SE to MS1B medium and culture

under light (65 μM m

−2 s

−1 and 16 h photoperiod) at 26 °C for

embryo germination ( see Note 17 ).

2. After 3 weeks, transfer well-developed plants with a robust

shoot and root system to plastic pots containing sterile Pro-

Mix BX potting mix (Premier Horticulture Inc., Red Hill, PA)

and acclimate in a growth room in light (65 μM m

−2 s

−1 and

16-h photoperiod) at 26 °C.

3. After 4 weeks, transfer well acclimated, vigorously growing

plants to a greenhouse.

4. Conﬁ rm gene expression in regenerated plants by observing

various plant tissues using a stereomicroscope equipped for

epi- ﬂ uorescence ( see Note 18 ).

4 Notes

1. Dormant cuttings are obtained by pruning annual wood from

grapevines during the winter season. Alternatively, certiﬁ ed

cuttings can be obtained from grapevine germplasm reposito-

ries such as the University of California, Davis, or the USDA

cold-hardy grapevine repository in Geneva, NY.

2. An embryogenic response from unopened leaf explants on

NB2 medium is observed from all seedless cultivars tested,

whereas a majority of seeded cultivars will only respond using

the stamen/pistil procedure. This factor must be considered

prior to embryogenic culture initiation from leaf explants.

3. Production of embryogenic responses from stamen and pistil

explants varies widely with Vitis species and cultivar. Hence

untested individual cultivars must be evaluated on each induc-

tion medium listed above to obtain an embryogenic response.

A list of responsive varieties and their optimum induction

media can be found in our reference publication [

13 ].

4. The use of TC agar (Phytotechnology Laboratories, LLC,

Shawnee Mission, KS, USA, Catalog No. A 175) or an agar

brand of equivalent purity, is paramount for successful induc-

tion and maintenance of embryogenic cultures. Use of other

gelling agents in X6 medium results in a rapid decline in

embryogenic potential and eventual culture death. A simple

observation to judge agar purity is the relative translucence of

poured dishes: the more translucent, the better.

5. To make a stock solution of Fe–EDTA, dissolve 7.44 g of

Na

2 EDTA·2H

2 O and 1.86 g FeSO

4 ·7H

2 O in sterile distilled

water and make ﬁ nal volume to 1 L. Although a number of

bacterial media were used for Agrobacterium culture, MG/L

medium provides better cell quality by avoiding overgrowth

and assists in maintaining bacterial virulence.

3.7 Somatic Embryo

Germination and Plant

Regeneration

Somatic Embryogenesis in Vitis

274

6. Rifampicin is dissolved in methanol or DMSO for making

stock solutions. Carbenicillin , cefotaxime and kanamycin sul-

fate are dissolved in distilled water and then ﬁ lter sterilized. All

antibiotic solutions are stored at −20 °C and thawed just prior

to use. Antibiotics are added to culture medium after auto-

claving and cooling the medium to 55 °C. Rifampicin is light

labile; preparation of stock solutions must be accomplished in

very dim light with storage in the dark.

7. Micropropagation cultures are initiated from the earliest

sprouting shoots of previously dormant ﬁ eld-grown plants

when they reach approximately 10 cm in length. Micro-

dissected shoot apical meristems are used as explants. We pre-

viously determined that shoot apical meristems taken at this

stage and from the ﬁ eld consistently yield the most sterile and

vigorous micropropagation cultures. Cultures are initiated on

C2D4B medium, with ﬁ ve meristems per dish [

25 ] and are

ultimately used for obtaining unopened leaf explants.

8. It is critical to use only unopened leaves of speciﬁ c size. Use of

larger leaf explants will produce solely non-embryogenic cul-

tures with no regeneration ability.

9. Stamen and pistil explants can be divided into 4 developmen-

tal stages based on size of inﬂ orescences and individual ﬂ ow-

ers, anther size and anther color. Stage II and III explants are

known to produce an embryogenic cultures in a large percent-

age of cultivars tested [

12 ].

10. It is critical to carefully excise intact stamens (anther with

attached ﬁ lament) and place all stamens from ﬁ ve ﬂ owers in a

clump/group to obtain an optimal embryogenic response. No

embryogenic response will be obtained with damaged or

detached ﬁ laments.

11. Embryogenic response from stamen and pistil explants is gen-

otype dependent. In general, a greater number of cultivars

produce an embryogenic response from stamen explants [

13 ].

12. A difference in culture proliferation rates and persistence is

observed among various cultivars in both solid and liquid

medium. This factor must be considered to ensure transfer

to fresh medium at the right interval and avoid culture

browning.

13. The use of a stereomicroscope in order to select proper tissue

for transfer is an absolute requirement to accomplish this pro-

cedure (Fig.

3b ) and cannot be stressed enough. It is impor-

tant to selectively transfer rapidly proliferating PEM to fresh

X6 medium using a microscope at 4–6-week interval (Fig.

3c,

e ). Failure to do so will lead to asynchrony of cultures, preco-

cious SE germination , decrease in embryogenic competence

and eventual termination of cultures.

Sadanand A. Dhekney et al.

275

14. It is important to use rapidly growing embryogenic cultures

for gene insertion. Use of older cultures can result in signiﬁ -

cantly lower-to-none insertion frequency and poor plant

regeneration.

15. Cocultivation of SE on ﬁ lter paper dramatically improves gene

insertion efﬁ ciency while preventing bacterial overgrowth and

culture necrosis [

19 ].

16. Proliferation of grapevine embryogenic cultures occurs by

direct secondary embryogenesis with new embryos arising

from the surface cells of existing SE or pre-existing embryo-

genic calli (Fig.

3f ). Thus, surface cells of cotyledonary-stage

SE are ideal targets for gene insertion and plant regeneration.

17. Plant recovery from germinated somatic embryo s can be

enhanced by trimming enlarged, ﬂ eshy cotyledons. This

response is species and cultivar dependent and needs to be

tested for speciﬁ c cultivars [

16 ]. A newly published two-step

culture procedure dramatically improves plant recovery [

26 ].

This includes culturing embryos on C2D4B medium for a

3-week period followed by transferring the germinated

embryos to MSN medium.

18. Uniform GFP expression is observed in plant tissues including

leaves, roots, ﬂ owers, stamens, and pistils (Fig.

4 ). Gene insertion

efﬁ ciency varies widely with Vitis species and cultivars [

16 ].

Fig. 4 GFP expression in a genetically modiﬁ ed grapevine . Uniform expression is observed in somatic embryo s

( a ), leaves and tendrils ( b ), roots ( c ), inﬂ orescences ( d ), stamens, and pistils ( e ). Note that the central glowing

spot in ( e ) represents the stigma (reproduced from Refs. 18 and 15 with permission from Springer)

Somatic Embryogenesis in Vitis

276

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Among the various species and cultivars tested, “Thompson

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produces the highest number of modiﬁ ed embryo and plant

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ﬂ oral organs. Thus, it is an ideal model with which to learn the

procedures.

Acknowledgements

S.A. Dhekney holds the E.A. Whitney Endowed Professorship in

the UW Department of Plant Sciences. The research that enabled

the Precision Breeding approach for grapevine was fostered by

long-term support from the Florida Department of Agriculture

and Consumer Services Viticulture Trust Fund, the USDA

Specialty Crops Research Initiative Grant Program, and the Florida

Agricultural Experiment Station, UF/IFAS.

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vol. 1359, DOI 10.1007/978-1-4939-3061-6_12, © Springer Science+Business Media New York 2016

Chapter 12

Somatic Embryogenesis in Peach-Palm ( Bactris gasipaes )

Using Different Explant Sources

Douglas A. Steinmacher , Angelo Schuabb Heringer , Víctor M. Jiménez ,

Marguerite G. G. Quoirin , and Miguel P. Guerra

Abstract

Peach palm ( Bactris gasipaes Kunth) is a member of the family Arecaceae and is a multipurpose but

underutilized species. Nowadays, fruit production for subsistence and local markets, and heart-of-palm

production for local, national, and international markets are the most important uses of this plant.

Conventional breeding programs in peach palm are long-term efforts due to the prolonged generation

time, large plant size, difﬁ culties with controlled pollination and other factors. Although it is a caespitose

palm, its propagation is currently based on seeds, as off-shoots are difﬁ cult to root. Hence, tissue culture

techniques are considered to be the most likely strategy for efﬁ cient clonal plantlet regeneration of this

species. Among various techniques, somatic embryogenesis offers the advantages of potential automated

large-scale production and putative genetic stability of the regenerated plantlets. The induction of somatic

embryogenesis in peach palm can be achieved by using different explant sources including zygotic embryos,

immature inﬂ orescences and thin cell layers from the young leaves and shoot meristems. The choice of

a particular explant depends on whether clonal propagation is desired or not, as well as on the plant

conditions and availability of explants. Protocols to induce and express somatic embryogenesis from

different peach palm explants, up to acclimatization of plantlets, are described in this chapter.

Key words Clonal propagation , Conservation programs , Heart-of-palm , Large-scale production ,

Pejibaye palm , Somatic embryo

1 Introduction

The peach palm ( Bactris gasipaes Kunth, Arecaceae) is a Neotropical

palm whose origin is still uncertain. Authors hypothesizing a sin-

gle origin point out to the western Amazon Basin, while those

supporting a multiple origin suggest the western and northwest-

ern sides of the Andes and lower Central America, in addition to

the western Amazon Basin, to be the centers of origin [

1 ]. It is

considered a multipurpose tree and plays an important role in

agroforestry in several Latin American countries [

2 ]. The produc-

tion of fruits and heart-of-palm for the national markets is one of

its most important uses, becoming peach palm the main source for

280

cultivated heart-of-palm [ 3 ]. Historically, this species has been

considered recalcitrant to in vitro culture. During the last few

years, however, several advances have been achieved with the suc-

cessful application of these techniques in this species; nevertheless,

a commercial protocol does not exist yet [

4 ].

Conventional breeding programs of peach palm are long-term

efforts due to long generation times of at least 6 years, large plant

size, difﬁ culties with controlled pollination, and other factors.

Therefore, in vitro clonal propagation has the potential to reduce

the time necessary for establishing elite plant orchards by capturing

and ﬁ xing the genetic gain expressed by selected plants for breeding

purposes. Peach palm conservation programs may also proﬁ t from

the use of in vitro regeneration protocols since germplasm banks

could be cloned and backups kept in other institutions for safekeep-

ing. Furthermore, somatic embryo genesis has the possibility to be

coupled to conservation programs through, for instance, the cryo-

preservation of somatic embryos [

5 ], as well as the production of

synthetic seeds for plantlet exchange. Hence, a reliable in vitro

regeneration protocol for peach palm is important and the develop-

ment of efﬁ cient methodologies is considered a necessity to support

use, conservation and breeding programs of this species [

1 ]. Among

the available techniques for in vitro plant generation, somatic

embryogenesis offers advantages such as large scale automated pro-

duction, cycling cultures through secondary embryogenesis [

6 ] and

genetic stability of the regenerated plantlets [

4 ]. For palm species,

somatic embryogenesis has been considered the preferred in vitro

regenerative pathway because of the larger number of regenerated

plantlets that can be produced compared to organogenesis [

7 – 9 ].

Tissue culture of palms is generally time consuming and the

biological events in each step of the process progress very slowly [

4 ,

10 ]. In peach palm, our experience shows that in vitro regeneration

of a reasonable number of plantlets takes about 2 years and one of

the critical aspects regarding the protocol is the choice of the

explants. Successful induction of somatic embryo genesis has already

been reported from different tissues, such as leaf primordia from

adult plants [

11 ], shoots and leaf primordia from in vitro- grown

plantlets [

9 ], immature inﬂ orescences [ 4 ], mature zygotic embryo s

[

12 ] and immature zygotic embryos [ 13 ]. The choice of the explant

source in this species depends also upon the aim pursued and

explant availability. For instance, the use of zygotic embryos as

explants might have limited applications in conservation programs;

however, they may serve as an interesting model to study peach

palm somatic embryogenesis because a relatively high induction

rate has been observed within few months of culture [

12 ] and the

morpho-histological responses from zygotic embryos were very

similar to those observed from shoot meristems and leaf sheaths

[

9 , 12 ]. However, for the clonal propagation and conservation of

selected genotypes, the development of protocols that allow regen-

eration from explants obtained from adult plants is necessary.

Douglas A. Steinmacher et al.

281

2 Materials

1. Zygotic embryo s as explants: Seeds from mature fruits, about

4 months after pollination, from one selected open pollinated

palm (Fig.

1 ) ( see Note 1 ).

2. Inﬂ orescences as explants: Immature inﬂ orescences from open

pollinated plants ( see Note 2 ).

3. 70 % (v/v) ethanol.

4. 40 % bleach solution containing at least 5 % of available chlo-

rine with one drop of the surfactant Tween 20

® for each

100 mL.

5. Sterile distilled water

6. Culture tubes (10 × 25 mm).

7. Disposal Petri dishes (90 × 15 mm).

8. Basal culture medium containing MS salts [

14 ], Morel vita-

mins [

15 ], 3 % (w/v) sucrose , 500 mg/L glutamine , with pH

adjusted to 5.8 prior to adding the gelling agent and auto-

claved for 15 min at 1 kgf cm

−2 .

9. Pretreatment liquid culture medium based on basal culture

medium supplemented with 1.5 g/L activated charcoal and

200 μM 2,4-dichlorophenoxyacetic acid (2,4-D).

10. Induction culture medium I based on basal culture medium

gelled with 2.5 g/L Gelrite and enriched with 1 μM AgNO

3 and

10 μM Picloram (4-amino-3,5,6-trichloropicolinic acid) [

12 ].

11. Induction culture medium II based on basal culture medium

supplemented with 1.5 g/L activated charcoal and 300 μM

Picloram gelled with 2.5 g/L Gelrite .

12. Growth culture medium based on basal culture medium sup-

plemented with 1.5 g/L activated charcoal and gelled with

7 g/L Agar .

Fig. 1 Fruits collected during ripening used as explant source. ( left ) Ripening fruits, showing characteristic

color. ( middle ) Seeds inside the fruits. ( right ) Mature and well-developed zygotic embryo used as explant for

induction of somatic embryo genesis

Somatic Embryogenesis in Peach-Palm

282

13. Maturation culture medium based on basal medium

supplemented with 2,4-D (40 μM), 2-isopentenyl adenine

(2-iP, 10 μM), activated charcoal (1.5 g/L), and increased

glutamine (1 g/L) plus hydrolyzed casein (0.5 g/L) as organic

nitrogen source and gelled with 2.5 g/L Gelrite .

14. Conversion culture medium constituted by basal media con-

taining 24.6 μM of 2-iP plus 0.44 μM of naphthalene acetic

acid and gelled with with 2.5 g/L Gelrite .

15. Expanded polystyrene trays, containing 5 × 5-cm cells.

16. Commercial substrate (e.g., PlantMax

® Paulinia, SP, Brazil,

electrical conductivity 1.5–2.0 dS/m) and carbonized rice

straw (1:1)

3 Methods

1. Remove the hard endocarp of the seeds to obtain the kernels

(i.e., zygotic embryo s enclosed in the endosperm ). Endocarp

can be easily removed without damaging the embryo by allow-

ing the former to dry slightly. Afterwards, let the seeds to

rehydrate in distilled water for further use.

2. Wash the kernels with running tap water and surface-sterilize

the hard endosperm with enclosed embryo by 1 min immer-

sion in 70 % ethanol, followed by 40-min soaking in the 40 %

bleach [

12 ].

3. Rinse the kernels with distilled-autoclaved water for at least

three times in the transfer hood.

4. Remove the zygotic embryo s (Fig.

2a ) from the endosperm in

the transfer hood with the help of a stereoscope.

5. Transfer the embryos to the induction culture medium I in

Petri dish.

6. After 1–2 weeks in culture, a swelling in the mesocotyl region

of the zygotic embryo will be observed. Histological analysis

has indicated that mitotic events occur in the subepidermic

tissue, mainly in cells adjacent to vascular bundles in the meso-

cotyl of the zygotic embryo [

12 ]. After 4 weeks, intense cel-

lular proliferation occurs in the cotyledonary blade showing

the ﬁ rst globular structures onto the primary callus (Fig.

2b ).

These initial globular structures will further develop into small

clusters of somatic embryo s (Fig.

2c ).

7. After 3 months in culture, up to 27 % of primary calli develop

embryogenic callus using this method [

12 ] (Fig. 2d ).

8. All steps of induction and expression of somatic embryo gen-

esis are to be undertaken in the dark in a growth chamber at

25 ± 2 °C.

3.1 Somatic

Embryogenesis

from Zygotic Embryos

Douglas A. Steinmacher et al.

283

1. Collect inﬂ orescences in the stage described above

(Subheading

2 ) from adult plants. Care should be taken to

avoid damaging the mother plant or the inﬂ orescence. The

inﬂ orescences must be promptly transported to the laboratory.

2. Remove the external spathes. Surface-sterilize the inﬂ ores-

cences when they are still surrounded by the internal spathes

by immersion in 70 % ethanol for 5 min, following by air-

drying in aseptic conditions.

3. Remove the internal spathes (with a size of approximately

5–8 cm) obtaining the explant that is going to be dissected

(Fig.

3a ). Separate the isolated inﬂ orescences in individual

rachillae and use them as explants ( see Note 3 ).

4. Place the dissected rachillae in culture tubes containing 25 mL

of pre-treatment liquid culture medium for 4 weeks with occa-

sional agitation.

3.2 Somatic

Embryogenesis

from Inﬂ orescences

Fig. 2 Induction of somatic embryo genesis from peach palm zygotic embryo s. ( a ) Zygotic embryo s used as

primary explant source. ( b ) Initial development of callus showing the initial development of globular structures

( arrow ). ( c ) Further development of the globular structures into small clusters after 6-week culture. ( d )

Development of a cluster of somatic embryos ( arrows ) on the callus after 12 weeks of culture (reproduced

from Ref. 17 with permission from Oxford University Press)

Somatic Embryogenesis in Peach-Palm

284

5. Afterwards dissect the rachillae into slices 1–2 mm thick and

inoculate them into Petri dishes containing induction culture

medium II.

6. Distinct in vitro responses might be observed, including oxi-

dation of the explants, development of ﬂ ower buds (Fig.

3b ),

dedifferentiation into actively growing tissue (Fig.

3c ), and

development of clusters of somatic embryo s (Fig.

3d ).

7. Up to 8 % of the explants can develop embryogenic callus after

32 weeks without subculturing [

4 ].

8. All steps of induction and expression of somatic embryo gen-

esis are to be undertaken in the dark in a growth chamber at

25 ± 2 °C.

Fig. 3 Somatic embryo genesis and plantlet regeneration from immature inﬂ orescences of peach palm. ( a )

Immature inﬂ orescences utilized as explant source (bar, 1 cm). ( b ) In vitro development of ﬂ ower bud ( arrow )

(bar, 1 mm). ( c ) Non-organized cellular proliferation of explants (bar, 2.5 mm). ( d ) Somatic embryogenic induc-

tion: note the development of globular somatic embryo s ( thin arrow ) and nodular tissue ( thick arrow ) (bar,

2.5 mm) (reproduced from Ref. 4 with permission from Springer Science and Business Media)

Douglas A. Steinmacher et al.

285

1. Remove the zygotic embryo s as indicated in Subheading 3.1

and transfer them to culture tubes containing 10 mL growth

culture medium. Keep cultures at 26 ± 1 °C in a 16-h photo-

period, with 50–60 μmol m

−2 s

−1 provided by cool-white ﬂ uo-

rescent lamps, until the plantlets reach 5–8 cm in height.

2. Remove the leaves, roots, any haustorial tissue and the most

external green leaf sheath of the plantlets. Section the remain-

ing embryo axis transversely in 0.7–1.0 mm slices to obtain

different histogenic layers (Fig.

4a ) ( see Note 4 ).

3. Inoculate the thin cell layer s in Petri dishes containing induc-

tion culture medium II.

4. Subculture to the same culture medium only after develop-

ment of callus is evident. Using this procedure, up to 43 % of

the explants can develop embryogenic callus [

10 ].

5. All steps for induction and expression of somatic embryo genesis

are to be undertaken in the dark in a growth chamber at 25 ± 2 °C.

3.3 Use of Thin Cell

Layers as Explants

to Induce Somatic

Embryogenesis

Fig. 4 Schematic diagram showing the origin of the explants utilized for the thin cell layer method ( dark scale

bar , 1.75 cm; white scale bar , 3 mm)

Somatic Embryogenesis in Peach-Palm

286

1. Embryogenic calli developed from any of the protocols

described above are similar and result in the development of

clusters of somatic embryo s; therefore, all might be further

cultured following the next steps.

2. Subdivide embryo clusters into smaller clusters of 5–8 somatic

embryo s before transferring to maturation conditions.

3. Transfer the embryo clusters to maturation culture medium

and incubate under dark conditions.

4. These cultures are subculture every 4 weeks in new fresh cul-

ture medium.

5. To convert mature somatic embryo s into plantlets, transfer them

to conversion culture medium in Petri dishes. Keep the cultures

at 25 ± 2 °C under a 16-h photoperiod (50–60 μmol m

−2 s

−1 pro-

vided by cool-white ﬂ uorescent lamps) for 4 weeks.

6. Subsequently, transfer the plantlets to growth culture medium

until they are 6 cm tall, when they can be acclimatized.

7. For acclimatization remove plantlets from the culture vessel,

wash all culture medium remnants thoughtfully and prune the

root system to approximately 2 cm (Fig.

5a ). Transfer the plant-

lets to a commercial substrate in expanded polystyrene trays.

8. Allocate the trays inside a plastic box covered with glass

(Fig.

5b ) to allow the entry of light and reduce water loss.

Keep these plantlets under 16-h photoperiods with 100–

130 μmol m

−2 s

−1 light intensity provided by ﬂ uorescent and

sodium vapor lamps in a growth chamber.

9. After 4 weeks start gradually opening the glass cover to increase

gas exchange and reduce relative humidity.

10. Transfer the plantlets to plastic bags containing the same sub-

strate and move them to the greenhouse with shading.

11. This acclimatization system presented high survival

(84.2 ± 6.4 %) after 16 week (Fig.

5c ) [ 13 ] .

3.4 Conversion

of Somatic Embryos

and Plantlet

Acclimatization

Fig. 5 Acclimatization of peach palm plantlets. ( a ) Plantlets used for acclimatization and with pruned roots

(bar, 2.5 cm). ( b ) Apparatus utilized for acclimatization (bar, 12.5 cm). ( c ) Acclimatized plantlets (bar, 5 cm)

(reproduced from Ref. 12 with permission from Springer Science and Business Media)

Douglas A. Steinmacher et al.

287

4 Notes

1. Our practical experience suggests that most appropriate stage

is when fruits are changing from green color to its characteris-

tic ripe color (yellow to red). During this stage, low contami-

nation rate are observed and the explants respond promptly to

the in vitro conditions.

2. The inﬂ orescences’ developmental stage on SE, these were

classiﬁ ed as Inﬂ 1, Inﬂ 2, and Inﬂ 3, according to the external

spathes’ size from 5 to 8, 8 to 12, and 12 to 16 cm, respec-

tively. According to [

16 ], these inﬂ orescences are formed in

the axils of leaves 2–5, 6–9, and 10–15, respectively, where leaf

1 is the newest expanded leaf in the crown.

3. The rachis is naturally sterile inside when the internal spate is

intact and healthy. If the rachis appears to be oxidized or con-

taminated, low success rate is observed.

4. This explant source might also be obtained from off-shoots of

adult selected palm trees.

Acknowledgements

The authors thank the Coordenação de Aperfeiçoamento de

Pessoal de Nível Superior—CAPES, Ministry of Education

(Brasília, Brazil), the Conselho Nacional de Desenvolvimento

Cientíﬁ co e Tecnológico—CNPq (Brasília, Brazil), Fundação

Araucária, (Curitiba, PR, Brazil), Parque Tecnológico da Itaipu- PTI

(Foz do Iguaçu, PR, Brazil), and Consejo Nacional para

Investigaciones Cientíﬁ cas y Tecnológicas (San José, Costa Rica),

for their support.

References

1. Mora-Urpí J, Weber JC, Clement CR (1997)

Peach palm ( Bactris gasipaes Kunth). Institute

of Plant Genetics and Crop Plant Research and

International Plant Genetic Resources

Institute, Rome

2. Clement CR, Mora-Urpí JE (1987) Pejibaye

palm ( Bactris gasipaes , Arecaceae): multi-use

potential for the lowland humid tropics. Econ

Bot 41:302–311

3. Clement CR (2008) Peach palm ( Bactris gasi-

paes ). In: Janick J, Paull RE (eds) The encyclo-

pedia of fruit and nuts. CABI, Wallingford,

UK, pp 93–101

4. Steinmacher DA, Clement CR, Guerra MP

(2007) Somatic embryogenesis from immature

peach palm inﬂ orescence explants: towards

development of an efﬁ cient protocol. Plant

Cell Tissue Organ Cult 89:15–22

5. Heringer AS, Steinmacher DA, Fraga HPF,

Vieira LN, Ree JF, Guerra MP (2013) Global

DNA methylation proﬁ les of somatic embryos

of peach palm ( Bactris gasipaes Kunth) are

inﬂ uenced by cryoprotectants and droplet-

vitriﬁ cation cryopreservation. Plant Cell Tissue

Organ Cult 114:365–372

6. Perez-Nunez MT, Chan JL, Saenz L, Gonzalez

T, Verdeil JL, Oropeza C (2006) Improved

somatic embryogenesis from Cocos nucifera

(L.) plumule explants. In Vitro Cell Dev Biol

Plant 42:37–43

7. Guerra MP, Torres AC, Teixeira JB (1999)

Embriogênese somática e sementes sintéticas.

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Cultura de tecidos e transformação genética

de plantas. SPI/EMBRAPA, Brasilia, Brazil,

pp 533–568

8. Sluis CJ (2006) Integrating automation tech-

nologies with commercial micropropagation.

In: Dutta Gupta S, Ibaraki Y (eds) Plant tissue

culture engineering. Springer, The

Netherlands, pp 231–251

9. Steinmacher DA, Krohn NG, Dantas ACM,

Stefenon VM, Clement CR, Guerra MP

(2007) Somatic embryogenesis in peach palm

using the thin cell layer technique: induction,

morpho- histological aspects and AFLP analysis

of somaclonal variation. Ann Bot 100:

699–709

10. Verdeil JL, Huet C, Grosdemange F, Buffard-

Morel J (1994) Plant regeneration from cul-

tured immature inﬂ orescences of coconut

( Cocos nucifera L)—evidence for somatic

embryogenesis. Plant Cell Rep 13:218–221

11. Almeida M, Almeida CV (2006) Somatic

embryogenesis and in vitro plant regeneration

from pejibaye adult plant leaf primordial. Pesq

Agrop Brasileira 41:1449–1452

12. Steinmacher DA, Cangahuala-Inocente GC,

Clement CR, Guerra MP (2007) Somatic

embryogenesis from peach palm zygotic

embryos. In Vitro Cell Dev Biol Plant 43:

124–132

13. Maciel SA, Fermino Junior PCP, da Silva RA,

Scherwinski-Pereira JE (2010) Morpho-

anatomical characterization of embryogenic

calluses from immature zygotic embryo of

peach palm during somatic embryogenesis.

Acta Sci Agron 32:263–267

14. Murashige T, Skoog F (1962) A revised medium

for rapid growth and bio assays with tobacco

tissue cultures. Physiol Plant 15:473–497

15. Morel G, Wetmore RH (1951) Fern callus tis-

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16. Clement CR (1987) Preliminary observation

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( Bactris gasipaes H.B.K.) inﬂ orescences. Rev

Biol Trop 35(1):151–153

17. Steinmacher DA, Guerra MP, Saare-Surminski

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_13, © Springer Science+Business Media New York 2016

Chapter 13

Somatic Embryogenesis: Still a Relevant Technique

in Citrus Improvement

Ahmad A. Omar , Manjul Dutt , Frederick G. Gmitter , and Jude W. Grosser

Abstract

The genus Citrus contains numerous fresh and processed fruit cultivars that are economically important

worldwide. New cultivars are needed to battle industry threatening diseases and to create new marketing

opportunities. Citrus improvement by conventional methods alone has many limitations that can be over-

come by applications of emerging biotechnologies, generally requiring cell to plant regeneration. Many

citrus genotypes are amenable to somatic embryogenesis, which became a key regeneration pathway in

many experimental approaches to cultivar improvement. This chapter provides a brief history of plant

somatic embryogenesis with focus on citrus, followed by a discussion of proven applications in

biotechnology- facilitated citrus improvement techniques, such as somatic hybridization, somatic cybrid-

ization, genetic transformation, and the exploitation of somaclonal variation. Finally, two important new

protocols that feature plant regeneration via somatic embryogenesis are provided: protoplast transforma-

tion and Agrobacterium -mediated transformation of embryogenic cell suspension cultures.

Key words Agrobacterium -mediated transformation , Cell suspension , Cybridization , Polyethylene

glycol ( PEG ) , Protoplast fusion , Protoplast transformation , Somaclonal variation , Somatic hybrid

1 Introduction

Citrus spp., native of South East Asia and China, are cultivated in

more than 100 countries, between approximately 40° N and 40° S

around the world. The genus Citrus has been recognized as one of

the most economically important fruit tree crops in the world. The

most commercially important Citrus species are oranges ( Citrus

sinensis L. Osbeck), tangerines ( Citrus unshiu Marc., Citrus nobilis

Lour., Citrus deliciosa Ten., Citrus reticulata Blanco and their

hybrids), lemons ( Citrus limon L. Burm. f.), limes ( Citrus auran-

tifolia Christm. Swing. and Citrus latifolia Tan.), and grapefruits

( Citrus paradisi Macf.). Fortunella , Poncirus , Microcitrus ,

Clymenia , and Eremocitrus are other genera of the family Rutaceae ,

related to Citrus . The importance of Citrus spp. is linked to their

economic value and to the nutritional proprieties of their fruits.

290

Moreover, Citrus spp. are connected to the social background of

the countries where they are grown, because many traditions, also

those related to the cookery, involve the use of Citrus fruits. Citrus

fruits are mostly eaten fresh, but a large part of the production,

mainly of grapefruits and oranges, is also used for juice extraction.

Furthermore, Citrus spp. are utilized in several ﬁ elds, not only in

the food industry, such as the production of marmalades, candies,

etc., but also, due to their richness in essential oils and polyphe-

nols, in the cosmetic, ﬂ avor, and pharmacy industries.

Although a high genetic variability is present in the genus

Citrus and its wild relatives, improvement by conventional breed-

ing is difﬁ cult because of various biological factors including ste-

rility [

1 ] self- and cross-incompatibility [ 2 ], widespread nucellar

embryony [

1 , 3 ], and long juvenile periods resulting in large plant

size at maturity. A consequence of these factors is the dearth of

information on genetic control of economically important traits

and rapid and effective screening procedures [

4 ]. Sweet orange

and grapefruit are important citrus species, and they are believed

to be interspeciﬁ c hybrids, not true biological species [

5 , 6 ]. All

cultivars within these species have arisen via somatic mutation,

either bud-sport or nucellar-seedling variants [

7 ], and not sexual

hybridization; intraspeciﬁ c hybridization results in weak or invia-

ble hybrid progeny (indicative of inbreeding depression) that gen-

erally produces fruit unlike those of the parents. The hybrid

orange cultivar Ambersweet, which originated by hybridization of

a mandarin × tangelo hybrid with sweet orange [

8 ], may be the

only exception.

Advances in in vitro tissue culture and improvements in molec-

ular techniques offer new opportunities for developing novel citrus

cultivars as some of these technologies can overcome the limita-

tions of sexual hybridization. For example, somatic hybrid ization

can create new combinations that were previously impossible

because of sterility or sexual incompatibility. By using this tech-

nique, improved varieties of citrus and unique new breeding par-

ents, for scion as well as for rootstocks, can be produced. This

technique consists of combining complementary parents with the

purpose of transferring desired traits to new plants such as resis-

tance to Phytophthora , citrus canker, citrus greening (HLB), citrus

variegated chlorosis, blight, and drought [

9 , 10 ]. Selecting somatic

mutations or genetic transformation allow the modiﬁ cation of very

few traits while retaining the essential characterization that typiﬁ es

speciﬁ c cultivar or cultivars groups. These techniques often require

somatic embryo genesis for efﬁ cient plant recovery. This chapter

will review somatic embryogenesis, and discuss applications of

in vitro biotechnologies and their protocols by utilizing somatic

embryogenesis in plant recovery, that can be used to obtain useful

new genetic combinations for citrus improvement.

Ahmad A. Omar et al.

291

Somatic embryo genesis is deﬁ ned as the differentiation of somatic

cells into somatic embryo s which show several distinct characteris-

tics [

11 ], including similarity to the developmental stages of

zygotic embryo genesis : bipolar structure presenting shoot and

root meristems, a closed tracheal system separated from the mater-

nal tissue, and frequently single-cell origin with production of spe-

ciﬁ c proteins. Somatic embryos play an important role in many

ﬁ elds, particularly for large-scale vegetative propagation. This mor-

phogenic process, that can occur with the formation of embryos

emerging directly from explants (direct somatic embryogenesis,

DSE), or after the formation of callus (indirect somatic embryo-

genesis, ISE), has been reported in several species [

12 , 13 ]. Somatic

embryos developing via DSE are formed from competent explant

cells which, contrary to ISE, are able to undergo embryogenesis

without dedifferentiation, i.e., callus formation. It is believed that

both processes are extremes of one continuous developmental

pathway [

14 ]. Distinguishing between DSE and ISE can be difﬁ -

cult [

15 ], and both processes have been observed to occur simul-

taneously under the same tissue culture conditions [

16 ]. Secondary

somatic embryos can arise cyclically from the surface of primary

somatic embryos, often at a much higher efﬁ ciency for many plant

species [

17 , 18 ]. Some cultures are able to retain their competence

for secondary embryogenesis for many years and thus provide use-

ful material for various studies, as described for Vitis ruperis [

19 ].

It is possible to induce somatic embryogenesis using different types

of culture media, environmental conditions and explants including

seedlings and their fragments, petioles, leaves, roots, shoot meri-

stems, seeds, cotyledons, anthers, pistils, and zygotic embryos.

Immature zygotic embryos present the most frequently applied

source of embryogenic cells which have been employed in most of

the established protocols. Immature zygotic embryos made possible

the induction of SE in plant species which, for many years, had been

considered to be recalcitrant, viz grasses [

20 ] and conifers [ 21 ].

By 1995 tissue culture conditions for SE induction had been

described for over 200 plant species [

17 ]; increasing numbers of

protocols were published after that. The most frequent mode of

embryogenesis is via callus formation, which is an indirect type of

regeneration.

The interest in somatic embryo genesis is due to several factors

such as high regeneration efﬁ ciency and the infrequent appearance

of somaclonal variation [

22 ]. Somatic embryo genesis has a key role

in in vitro clonal propagation for plant mass propagation, as well as

for germplasm conservation and exchange, cryopreservation to

establish gene banks, sanitation, metabolite production, and syn-

thetic seed production. The application of synthetic seed technol-

ogy to Citrus has been reported for somatic embryos of Citrus

reshni , Citrus reticulata Blanco (cv Avana and cv Mandarino

Tardivo di Ciaculli), Citrus clementina Hort. ex Tan. (cv Monreal

1.1 Somatic

Embryogenesis

in Plants

Somatic Embryogenesis in Citrus

292

and cv Nules), a lemon hybrid [ 23 , 24 ], and Kinnow mandarin [ 25 ].

Moreover, in vitro conservation of several Citrus species using

encapsulation –dehydration technology of cryopreservation has

also been reported [

26 , 27 ].

Plant regeneration systems that limit or avoid genetic chime-

rism in regenerants are of special value for biotechnologies that

combine tissue culture with genetic transformation or mutant

induction and selection. Genetic modiﬁ cation is a unicellular event,

and hence regeneration from multicellular centers frequently

results in the formation of genetic chimeras. A high probability for

the single cell origin of regenerants is what provides for ideal

SE. The classical conception of SE is based on the unicellular ori-

gin of somatic embryo s [

28 ], and this mode of somatic embryo

development was the most frequently noticed in embryogenic cell

suspension s of D. carota [

29 ]. However, single-cell origin of

somatic embryos is not the rule, and even in a model system such

as embryonic cell suspension of Daucus carota , development of

embryos from a group of cells cannot be excluded [

30 ].

Development of somatic embryos from more than one cell has in

fact been reported in several plant systems. Moreover, both a mul-

ticellular and a unicellular origin of somatic embryos in the same

regeneration system is quite a common phenomenon, as was

observed in several species including Musa spp. [

31 ], Cocos nucifera

[

32 ], Santalum album and S. spicatum [ 33 ], and H. vulgare [ 34 ].

It is believed that somatic embryos originated from a single cell

displayed normal morphology of “single embryo” while aberrant,

multiple embryos are derived from a group of cells [

35 – 37 ].

Numerous published protocols on successful SE induction and

plant regeneration in different plant species, suggest that SE could

be achieved for additional plant species provided that appropriate

explant and culture conditions are employed, although progress

will probably remain slow with the more recalcitrant woody

species.

The establishment of efﬁ cient embryogenic cultures has become

an integral part of plant biotechnology as regeneration of trans-

genic plants in most of the important crops (such as canola , cas-

sava, cereals, cotton, soybean, and various woody tree species) is

dependent on the formation of somatic embryo s. One of the most

attractive features of embryogenic cultures is that plants derived

from them are predominantly normal and devoid of any pheno-

typic or genotypic variation, possibly because they are often derived

from single cells and there is stringent selection during embryo-

genesis in favor of normal cells [

38 ]. Embryogenic cultures were

ﬁ rst described in callus and suspension cultures of carrot by Reinert

[

39 ] and Steward et al. [ 40 ], respectively. In the following decades

with increasing understanding of the physiological and genetic

regulation of zygotic as well as somatic embryogenesis, embryogenic

1.1.1 Somatic

Embryogenesis Protocol

Development

Ahmad A. Omar et al.

293

cultures had been obtained on chemically deﬁ ned media in a wide

variety of species [

38 ]. In most instances the herbicidal synthetic

auxin 2,4-dichlorophenoxyacetic acid (2,4-D) was required for the

initiation of embryogenic cultures; somatic embryos develop when

such cultures are transferred to media containing very low amounts

of 2,4-D or no 2,4-D at all.

During the 1950s a number of attempts were made to demon-

strate the totipotency of plant cells. The ﬁ rst evidence of the pos-

sibility that single cells of higher plants could be cultured in

isolation was provided by Muir et al. [

41 ], who obtained sustained

cell divisions in single cells of tobacco placed on a small square of

ﬁ lter paper resting on an actively growing callus, which served as a

nurse tissue. Similar results were obtained by Bergmann [

42 ] who

plated single cells and cell groups suspended in an agar medium.

Further progress was made by Jones et al. [

43 ], who were able to

culture single isolated cells in a conditioned medium in specially

designed microculture chambers. Direct and unequivocal evidence

of the totipotency of plant cells was ﬁ nally provided by Vasil and

Hildebrandt [

44 , 45 ], who regenerated ﬂ owering plants of tobacco

from isolated single cells cultured in microchambers, without the

aid of nurse cells or conditioned media. Up to date, in vitro culture

techniques have enabled plant regeneration from over 1000 differ-

ent species [

46 ], following two alternative morphogenetic path-

ways, shoot organogenesis (SO) or SE. Both morphogenic

pathways, SE and SO, may be induced simultaneously in the same

tissue culture conditions [

47 ]. Thus, differentiation between SE

and SO can sometimes be difﬁ cult, and even a detailed comparative

histological analysis of the morphogenic process can only suggest

an embryo-like origin of developing structures [

48 ]. However, SE

and SO can be separated in space and time [

49 , 50 ] with the use of

appropriate medium composition, mainly type or concentration of

plant growth regulator s (PGRs).

The application of in vitro systems based on SE for plant regen-

eration is determined not only by a high efﬁ ciency of somatic

embryo formation, but frequently depends on capacity of the

embryos for complete plant development. The process of develop-

mental changes, which a somatic embryo undergoes, is called

“ conversion ”, and it involves the formation of primary roots, a

shoot meristem with a leaf primordium and greening of hypocotyls

and cotyledons [

51 ]. In numerous systems, in spite of the high

number of somatic embryos produced, problems with a lack or a

low frequency of embryo conversion into plants has occurred. To

stimulate embryo conversion, and to improve the efﬁ ciency of

plant regeneration, a number of different strategies have been

tested. Gibberellic acid ( GA

3 ) is frequently employed in media

used for somatic embryo conversion. It should be stressed that in

some systems, abnormal morphology of somatic embryos did not

decrease the chances of development into normal plants [

52 – 54 ].

Somatic Embryogenesis in Citrus

294

In a plant seed the embryo is generally formed following the fusion

of gametes from two parents during fertilization . However, some

species form embryos in the seed without fertilization. This kind of

reproduction is termed apomixis by which somatic cell-derived

embryos develop in a seed. Apomixis is a fairly uncommon trait in

plants, but approximately 400 species exhibit this type of propaga-

tion in nature [

55 ]. Apomixis is classiﬁ ed into apospory, diplo-

spory, and adventitious embryony according to the developmental

process of somatic embryo(s). In apospory and diplospory, apo-

mictic embryo(s) develop megagametophytic structure without

meiotic reduction, which is widely observed in grass species. On

the other hand, in adventitious embryony as observed in citrus and

mango ( Mangifera indica L.), somatic embryos are directly initi-

ated from nucellar cells in ovule tissue [

56 ]. In citrus, polyembry-

ony, speciﬁ cally adventitious embryony, is a common reproductive

phenomenon. Some cultivars develop many embryos in a seed,

such as Satsuma mandarin ( Citrus unshiu ) and Ponkan ( Citrus

reticulata ) which form 20 or more embryos in a seed. In contrast,

monoembryonic cultivars (e.g., Clementine , Citrus clementina ,

and Kinokuni mandarin, Citrus kinokuni ) form only a single,

zygotically derived embryo in each seed [

57 ]. Apomixis has great

potential as a breeding technology because introduction of apo-

mixis into non-apomictic plants enables clonal propagation with

genetically true seeds from hybrids. The potential economic ben-

eﬁ t of incorporation of apomixis in rice was estimated to exceed

US $2.5 billion per annum [

58 ]. Because of its economic potential

as a breeding technology, genomics-based approaches have been

applied to identify the gene responsible for apomixes [

59 , 60 ].

Somatic embryo genesis is particularly attractive in citrus because

many cultivars and accessions have the capacity for nucellar embry-

ony [

61 ]. Somatic embryogenesis has been induced directly in cul-

tured nucelli [

62 ] and undeveloped ovules [ 63 , 64 ] or indirectly

via callus formation [

65 – 69 ]. Embryogenesis has also been induced

from endosperm -derived callus [

70 ], juice vesicles [ 71 ], anthers

[

72 , 73 ], and styles [ 74 – 77 ].

In order to apply the techniques of modern plant biotechnology

to citrus breeding, it is necessary to develop reliable and efﬁ cient

plant tissue culture procedures for plant regeneration (Fig.

1 ).

In citrus, the production of embryogenic callus lines have been

reported from the culture of excised nucelli [

78 ], abortive ovules

[

79 ], unfertilized ovules [ 80 ], undeveloped ovules [ 64 ], isolated

nucellar embryos [

81 ], Satsuma juice vesicles [ 71 ], anthers [ 82 ],

styles and stigmas of different species of citrus [

75 , 83 ], as well as

from leaves, epicotyls, cotyledons and root segments of in vitro

grown nucellar seedling of C. reticulata Blanco [

84 ]. The embryo-

genic potential of citrus varies with genotype and type of explant.

One important application of this technique is the production of

1.1.2 Somatic

Embryogenesis in Citrus

Ahmad A. Omar et al.

295

virus-free citrus plants through somatic embryo genesis from

undeveloped ovules of some citrus species [

79 , 85 ]. Somatic

embryo s, embryogenic callus and cell cultures recovered from

in vitro cultured ovules have also been used to develop cryopreser-

vation strategies for germplasm conservation [

86 ], to generate

somaclonal variation [

87 ], and for protoplast fusion technologies

to generate somatic hybrid s and cybrids [

4 , 9 , 88 , 89 ]. Many citrus

species are found responsive to culture on a basal medium supple-

mented with malt extract , but embryogenesis has been enhanced

by the addition of other growth substances.

Fig. 1 Somatic embryo genesis in citrus. ( a ) Citrus ovules, ( b ) ovule derived embryogenic callus, ( c ) embryogenic

callus, ( d ) embryogenic cell suspension cultures, ( e ) protoplast derived micro-calli, ( f ) callus derived somatic

embryo s, ( g , h ) small-medium somatic embryos on cellulose acetate ﬁ lter papers, ( i ) enlarged embryos on

EME- maltose medium, ( j ) enlarged embryos and shoots on 1500 medium, ( k ) small plantlet on B+ medium,

( l ) plantlets on rooting medium

Somatic Embryogenesis in Citrus

296

Anther culture is a commonly used method to produce

haploids and doubled-haploids in Citrus , as well as in other fruit

crops [

90 – 92 ]. Citrus anther culture produced also somatic

embryo - derived regenerants in C. aurantium [

82 , 93 ], C. sinensis ,

C. aurantifolia [

94 ], C. madurensis [ 95 ], C. reticulata [ 93 , 96 ],

Poncirus trifoliata , the hybrid No. 14 of C. ichangensis × C. reticulata

[

97 ] and C. paradisi . While the somatic embryogenesis capacity of

Citrus has been found to vary with the cultivar and type of explant,

regeneration methods that involve the use of embryogenic callus

of nucellar origin (polyembryonic types) generally provide the

best results. Unfortunately, these systems either fail or provide

only poor results with monoembryonic species that produce only

zygotic embryo s. Kobayashi et al. [

98 ] cultured the ovules of 23

monoembryonic cultivars and never obtained nucellar embryos.

The selection of elite citrus plants is essential for the development

of efﬁ cient systems of somatic embryo genesis. For these purposes,

explants should be collected from selected elite specimens that are

visibly free from any symptoms of disease, stress or spontaneous

mutations (i.e., variegated fruits and leaves, variation in color, size

and shape of fruits, and various other plant abnormalities). Carimi

[

99 ] addressed several points to bear in mind when deciding upon

the choice of explant, i.e., (1) callus formation appears to depend

on the status of the tissue, (2) callus initiation occurs more readily

in tissues that are still juvenile, and (3) explants must contain liv-

ing cells. When ﬂ oral tissues and fruits are old, chances of callus

and embryo formation from undeveloped ovules, stigma, or style

explants decrease. Stigma and styles derived from immature ﬂ ow-

ers and undeveloped ovules from unripe fruits have higher

embryogenic potential s, although embryogenic callus lines have

been successfully initiated also from the undeveloped ovules of

mature fruits.

The composition of the media used for in vitro regeneration of

citrus somatic embryo s is based on the inorganic salts recom-

mended by Murashige and Skoog [

100 ] and on the organic com-

pounds suggested by Murashige and Tucker [

101 ]. Sucrose

(50 g/L) is usually used as the carbon source. When needed,

growth regulators can be added directly to the medium before or

after autoclaving. The pH of the medium is generally adjusted to

5.8. Normally, 8 g/L agar is used to solidify media for citrus tissue

culture. After preparing the media, it could be stored at room tem-

perature for several weeks before use. Starrantino and Russo [

64 ]

ﬁ rst reported somatic embryogenesis from undeveloped ovule cul-

ture. The percentage of embryogenic explants ranges from 0 % to

70 %, depending on the genotype. As mentioned, this regeneration

procedure does not work with monoembryonic genotypes [

102 ]

(for more details about how to initiate somatic embryogenesis

including embryogenic callus and suspension lines from undeveloped

1.1.3 Source of Explants

to Initiate Somatic

Embryos in Citrus

1.1.4 Somatic Embryo

Induction and Growth

Media in Citrus

Ahmad A. Omar et al.

297

ovule culture, see [ 9 , 10 , 103 ]). EBA (MT basal medium plus

0.01 mg/L 2,4-D and 0.1 mg/L 6-BAP) and DOG (MT basal

medium plus 5 mg/L kinetin) media are often used for embryo-

genic callus induction [

103 ].

Somaclonal variation , first defined and reviewed by Larkin and

Scowcroft [

104 ], is a commonly observed phenomenon in cell and

tissue cultures of different species regardless of the regeneration

system used [

105 ]. This variation involves changes in both nuclear

and cytoplasmic genomes, and their character can be of genetic or

epigenetic nature [

22 ]. Mechanisms which determine somaclonal

variation [

106 – 108 ], as well as the advantages and drawbacks of

in vitro produced plant variants [

109 – 111 ], have been widely dis-

cussed. The identification of valuable somaclonal variants holds

great promise for cultivar improvement, especially for the citrus

species that are difficult to manipulate by sexual hybridization [

4 ].

Somaclonal variation has been observed in citrus plants regener-

ated from nucellar callus of monoembryonic “ Clementine ” man-

darin [

85 ]. Callus lines have been selected for salt tolerance [ 112 ,

113 ] and regenerated into plantlets; however, regenerated plantlets

lacked internodes and hence could not be propagated further [

114 ].

C. limon embryogenic culture lines resistant to “mal secco” toxin

were selected. These lines produced somatic embryo s, which

retained resistance to the toxin [

115 ]. “Femminello” lemon

somaclones have also been evaluated for tolerance to mal secco by

artificial inoculation [

116 ]. Somaclones of “Hamlin,” “Valencia,”

“Vernia,” and “OLL” (Orie Lee Late) sweet oranges have been

obtained via regeneration from callus, suspension cultures, and/or

protoplasts, obtained via somatic embryogenesis, in efforts to

improve processing and fresh market sweet oranges [

87 , 117 ].

Significant variation has been observed in fruit maturity date, juice

quality, seed content and clonal stability. The University of Florida,

Institute of Food and Agricultural Science (UF/IFAS), through

Florida Foundation Seed Producers (FFSP), has released several

improved sweet oranges regenerated using the somatic embryo-

genesis pathway, such as “Valencia protoclone SF14W-62”

(Valquarius

® -U.S. Patent PP21,535, selected for 6–8 weeks early

maturity date), “Valencia protoclones N7-3” (U.S. Patent

PP21,224 and T2-21, seedlessness and late maturity), “Hamlin

protoclone N13-32” (improved juice color), and somaclones

“OLL-4” and “OLL-8” (high yield and juice quality, clonal stability).

We are also evaluating several hundred lemon somaclones (derived

from multiple commercial lemon cultivars) for fruit rind oil con-

tent and seed content. We have identified several seedless soma-

clones and somaclones that consistently yield more oil per unit of

rind surface area (Gmitter, Grosser and Castle, unpublished data).

It is clear that useful genetic variation can be obtained from

large enough populations of somatic embryogenesis-regenerated

somaclones.

1.2 Applications

of SE in Cultivar

Improvement of Citrus

1.2.1 Generation

of Somaclonal Variation

Somatic Embryogenesis in Citrus

298

As mentioned above, new cultivars of sweet orange have been

developed from populations of plants regenerated from proto-

plasts via somatic embryo genesis (protoclones) (Fig.

1 ). In plant

tissue culture history, embryogenic cell culture and the develop-

ment of protoplast technologies that require plant recovery are

closely linked. Although progress in the development of proto-

plast technologies has been made in other woody tree species,

including the regeneration of somatic embryos from protoplasts

isolated from embryogenic cells of Pinus taeda and Picea glauca

[

118 – 120 ], citrus has been the true model system in this regard

primarily due to its robust ability for somatic embryogenesis.

The limited range of the explant source from which morphoge-

netically competent tissues can be obtained has limited success

with protoplast culture in other tree species. Methods for the iso-

lation and culture of Citrus protoplasts from embryogenic callus

and suspension cultures, and subsequent plant regeneration are

well developed [

9 , 10 , 89 , 103 , 121 – 123 ]. Protoplast fusion tech-

niques have been used to generate somatic hybrid plants from

more than 500 parental combinations, including more than 300

from our laboratory (for reviews, see ref. [

4 , 9 , 10 , 88 , 124 ]).

As a by-product of protoplast fusion , hundreds of diploid cybrid

citrus plants have also been regenerated via somatic embryogene-

sis [

125 , 126 ]. Protoplasts have also been proven to be very useful

in the genetic transformation of plants [

127 – 130 ], including eco-

nomically important cereals [

131 ]. Once again, citrus has led the

way with genetic transformation of protoplasts among woody

fruit trees, with transformed plant recovery due to robust somatic

embryogenesis [

129 , 132 ].

The complex 8P protoplast culture medium of Kao and Michayluk

[

133 ] has been used for successful protoplast culture and plant

regeneration from embryogenic cultures of several plant species.

The success of this complex medium is probably due to the appro-

priate concentrations of the multivitamin, organic acid, and sugar/

alcohol additives that are combined with the basal medium formu-

lation. These additives seem to provide additional buffering capac-

ity and reduce the environmental stress on protoplasts by providing

required metabolic intermediates needed to sustain adequate cell

viability and totipotency . However, optimal basal tissue culture

media have been developed for most plant genera, and an efﬁ cient

protoplast culture medium may be developed for a particular genus

by simply supplementing the optimal basal medium with 8P mul-

tivitamin, organic acid, and sugar/alcohol additives. This approach

has been successful for Trifolium [

134 , 135 ] and Citrus [ 9 ].

Reducing or eliminating the ammonia content of the basal medium

has also been useful. Most basal media contain high levels of

NH

4 NO

3 that can often be toxic to protoplasts. Glutamine or

Ca(NO

3 )

2 have been found to be good alternative sources of N in

1.2.2 Protoplast

Regeneration via Somatic

Embryogenesis

1.2.3 Protoplast Isolation

and Culture

Ahmad A. Omar et al.

299

embryogenic suspension culture and protoplast culture media, as

demonstrated in H+H suspension culture medium and BH3

protoplast medium of citrus [

9 ], as well as in Populus protoplast

media [

136 ]. Vardi et al. [ 137 ] reported the ﬁ rst example of suc-

cessful citrus protoplast isolation and culture, followed by callus

formation and embryo differentiation. Subsequently, numerous

Citrus species have been regenerated from protoplasts via somatic

embryo genesis [

124 ]. Ohgawara et al. [ 138 ] obtained for the ﬁ rst

time somatic hybrid s of citrus regenerated via somatic embryogen-

esis, involving Citrus ( C. sinensis and Poncirus trifoliata ). Citrus

protoplasts can be isolated from different sources including

embryogenic cells (cultured on either solid or liquid media), non-

embryogenic callus, and leaves. Embryogenic cell cultures (on

solid or liquid media) yield protoplasts with the best potential for

proliferation and embryo regeneration. Leaves are another often

utilized source for protoplast isolation in Citrus , because leaf

protoplasts are generally easy to isolate and large amounts of pro-

toplasts are produced; however, they are not totipotent and do not

develop into somatic embryos. In vitro cultured nucellar seedlings

are becoming more commonly used as a source of leaf material for

protoplast isolation, as this source eliminates the need for harsh

decontamination. Leaf protoplasts are often used in somatic fusions

with embryogenic culture protoplasts, where the latter provides

the capacity for somatic embryogenesis and plant recovery in

somatic hybrids and cybrids . Embryogenic callus or suspension

cultures used for protoplast isolation should be in the log phase of

growth at the time of isolation. Friable tissue with low starch con-

tent generally gives the best results. Citrus embryogenic cultures

often require continual subculturing for long periods before they

reach adequate friability and appropriate starch levels for proto-

plast manipulation. Transferring Citrus callus to glutamine -

containing media can sometimes reduce the starch content of cells

to appropriate levels for protoplast isolation [

9 , 10 , 103 ]. A proce-

dure for the induction of suspension cultures from embryogenic

calli has been previously described [

9 , 10 , 103 , 139 ]. Suspension

cultures offer several distinct advantages over stationary cultures,

especially when conducting multiple experiments requiring large

volumes of explant. Suspension cultures quickly generated needed

volumes of explant for multiple experiments, and rapidly growing

suspension cells have thinner cell wall s that are more amenable to

enzyme digestion. Combining an enzyme solution (generally con-

taining cellulose and macerase) with a complex protoplast culture

medium may reduce stress on protoplasts during isolation and

thereby increase viability. We prefer maintaining suspension cul-

tures on a 2-week subculture cycle, with optimum protoplast isola-

tions occurring at days 4–12, when suspension cultures are in the

log phase of growth.

Somatic Embryogenesis in Citrus

300

Somatic hybrid ization allows production of somatic hybrid s that

incorporate genomes of the two parents with little or no recombi-

nation, but with increased heterozygosity in the resulting poly-

ploidy hybrids [

140 ]. Somatic hybridization in citrus relies on the

process of somatic embryo genesis for plant regeneration. In citrus,

this technology has been extensively used and has important appli-

cations in both scion and rootstock improvement [

124 ]. The first

successful protoplast isolations were reported as early as 1982

[

123 ], and the first citrus somatic hybrid was obtained between

C. sinensis and P. trifoliata [

138 ]. These results encouraged the

development and incorporation of somatic hybridization tech-

niques into the citrus breeding programs in several countries [

9 ].

Somatic hybridization has made it possible to hybridize commercial

citrus with citrus relatives that possess valuable attributes, thus

broadening the germplasm base available for rootstock improve-

ment [

141 ]. Somatic hybrids have been developed and established

at the Citrus Research and Education Center, University of Florida,

USA for three decades to improve citrus scions and rootstocks [

9 ,

10 , 124 ]. The most important contribution somatic hybridization

can make to citrus breeding programs is the creation of unique

tetraploid breeding parents.

We have used somatic hybrid ization to create new tetraploid

somatic hybrids that combine elite diploid scion material as tetra-

ploid breeding parents being used in interploid hybridization

schemes to develop seedless and easy-to-peel new mandarin vari-

eties [

142 ], and in grapefruit/pummelo and acid fruit improve-

ment (lemons/limes) [

10 , 143 ]. The ﬁ rst seedless triploid mandarin

from this program (C4-15-19, from a cross of “LB8-9” with a

somatic hybrid of “Nova” mandarin hybrid + “Succari” sweet

orange), was recently released by UF/IFAS for commercialization.

This is the ﬁ rst released triploid citrus cultivar fathered by a

somatic hybrid. The majority of somatic hybrid breeding parents

produced for scion improvement have been from fusions of two

polyembryonic parents. In this case, the somatic hybrid can only

be efﬁ ciently used as a pollen parent in interploid crosses. Using

this approach, we have produced several thousand triploid hybrids

fathered by somatic hybrids. Interploid crosses utilizing a mono-

embryonic diploid female parent and a tetraploid male parent

require embryo rescue for triploid plant recovery because embryos

do not complete normal development, presumably as a conse-

quence of endosperm :embryo ploidy level balance. By contrast,

interploid crosses utilizing a monoembryonic tetraploid females

do not require embryo rescue [

10 ]. Somatic hybrid s produced

by the fusion of a polyembryonic embryogenic parent with a

monoembryonic leaf parent are frequently monoembryonic.

We have recently efﬁ ciently recovered triploid progeny by simply

planting fully developed seeds from interploid crosses involving the

1.2.4 Somatic

Hybridization

Scion Improvement

Ahmad A. Omar et al.

301

following monoembryonic somatic hybrid females in our breeding

program: “Succari” sweet orange + “Hirado Buntan” pummelo,

“Murcott” + “Chandler” sdlg.#80, “Murcott” + “Chandler” sdlg.

A-1-11 (grapefruit/pummelo improvement), “Santa Teresa”

lemon + “Lakeland Limequat” (lemon improvement), and

“W. Murcott” + UF03-B (“For tune” × “Murcott”) (mandarin

improvement) (J.W. Grosser, unpublished information). Thus, our

future somatic hybridization work will focus more on production

of monoembryonic somatic hybrids. Creation of triploid citrus

hybrids directly by electrofusion of haploid and diploid protoplasts

is also promising [

144 ].

Numerous allotetraploid somatic hybrid s via protoplast fusion

with plant recovery by somatic embryo genesis, which combine

complementary diploid rootstocks, have been produced [

9 ]. These

hybrids have direct rootstock potential [

145 ], but their most

important contribution may be their use as breeding parents in

rootstock crosses at the tetraploid level. We initiated tetraploid

rootstock breeding around the year 2000, and since this time hun-

dreds of zygotic allotetraploids (“tetrazygs”) have been obtained.

This approach is quite powerful genetically, because the alleles

from four rootstock genotypes can be recombined simultaneously,

creating a wealth of genetic diversity in progeny. Resulting allotet-

raploid rootstock candidates have been screened for tolerance to

the Diaprepes / Phytophthora complex [

117 , 87 ], salinity [ 145 ], and

now HLB (Huanglongbing or citrus greening), all with promising

results. With the cost of citrus production and harvesting increas-

ing over time, there has been greater emphasis on developing root-

stocks to facilitate Advanced Citrus Production Systems (ACPS),

that reduce tree size to make orchard management and crop har-

vesting more efﬁ cient and also to bring young trees into economi-

cally valuable production earlier. We learned early on that tetraploid

rootstocks, especially allotetraploid somatic hybrids, always have

some capacity to reduce tree size, even from somatic hybrids pro-

duced between two vigorous parents [

10 , 145 ]. Through multiple

ﬁ eld trials, we have identiﬁ ed some somatic hybrid and “tetrazyg”

rootstock hybrids that have combined desirable horticultural attri-

butes, disease resistance and stress tolerance traits, and confer vary-

ing degrees of tree size control [

10 ]. UF/IFAS has recently “fast

track” released 17 new rootstock selections to the Florida industry

for large scale evaluation that include one somatic hybrid and six

“tetrazyg” allotetraploid hybrids. The release additional improved

allotetraploid rootstocks can be expected in the near future.

Cybrids combine the nucleus of a species with alien cytoplas-

mic organelles [

126 , 146 ]. Cybridization could be a valuable

method for improvement of various crops that would be in the

non-regulated category of genetically modified organisms.

Rootstock Improvement

1.2.5 Somatic

Cybridization

Somatic Embryogenesis in Citrus

302

The first cybrids in citrus were created by the “donor–recipient”

method [

147 ]. The phenomenon of cybridization in citrus

also occurs as an accidental by-product of somatic hybrid iza-

tion via protoplast fusion [

125 , 148 ]. The general somatic

hybridization model of fusing embryogenic culture cell proto-

plasts with leaf protoplasts often yields diploid plants with the

morphology of the leaf parent. These plants have always, with-

out exception, been validated as cybrids, as citrus leaf proto-

plasts are not capable of plant regeneration. Such cybrids

always have the mitochondrial (mt) genome of the embryo-

genic suspension/callus parent, whereas the chloroplast (cp)

genome is randomly inherited. Thus, recovered cybrid plants

are regenerated via somatic embryo genesis. Moreira et al.

[

148 ] found that embryogenic suspension culture cells gener-

ally have four times more mt per cell than do leaf cells and

hypothesized that the extra mt acquired by the cybrid cells is

necessary to satisfy the high energy demand of the somatic

embryogenesis pathway of regeneration. This phenomenon

has been exploited to produce targeted cybrids. One approach

for cultivar improvement has been to transfer of cytoplasmic

male sterility (CMS) from “Satsuma” mandarin to other elite

but seedy scions via cybridization. This approach has the

potential to make existing popular cultivars less seedy, without

altering the cultivar integrity in any other way [

126 , 146 ].

This technique has only been partially successful in our experi-

ence; for example, we have produced cybrid “Sunburst” man-

darin clones that have less than half the normal seed content of

“Sunburst”, but still too many seeds to label them as seedless

(JW Grosser, unpublished information). However, these cybrid

“Sunburst” trees produce a fruit that is easier to peel and with

better flavor than traditional “Sunburst.” Accidental cybrids of

“Ruby Red” and “Duncan” grapefruit, both containing the mt

genome from “Dancy” mandarin, have also been produced

from separate experiments. In both cases, the fruit from cybrid

trees has improved characteristics, including significantly

higher brix and brix/acid ratios, and an extended harvesting

season that extends well into the summer with no vivipary or

fruit drying (Satpute et al., submitted). UF/IFAS has released

the first cybrid citrus cultivar, namely the N2-28 cybrid “Ruby

Red” grapefruit, from this work. We are also attempting to

utilize cybrid technology for improving disease resistance in

existing cultivars. The mt genome of kumquat ( Fortunella

crassifolia Swingle) is purported to contain a gene for citrus

canker resistance. Citrus canker disease has caused significant

damage to the Florida grapefruit industry. We have initiated an

embryogenic suspension culture of “Meiwa” kumquat and

performed fusions with leaf protoplasts of grapefruit cultivars

“White Marsh,” “Flame” (red) and a dark red somaclone N11-11.

Ahmad A. Omar et al.

303

Multiple diploid plants from each fusion combination exhibiting

grapefruit morphology have been regenerated and their cybrid

nature confirmed by mitochondrial intron marker analysis [

149 ].

Cp genome inheritance analysis in these plants is currently under-

way. These cybrid grapefruit plants are being propagated for a can-

ker challenge assay to determine if the transfer of the kumquat mt

genome can indeed improve their resistance to citrus canker.

Genetic transformation has become an attractive alternative

method for improving plant species including citrus, because it is

possible to maintain cultivar integrity while adding a single trait.

Exploiting the process of somatic embryo genesis, citrus can be

transformed either directly from embryogenic cell suspension

cultures or indirectly from isolated protoplasts. Embryogenic cells

are usually treated with an Agrobacterium culture followed by

selection and regeneration of transgenic plants. Plant protoplasts

are commonly transformed using the polyethylene glycol ( PEG )-

mediated DNA uptake process, and less frequently using electro-

poration. The PEG-mediated DNA transfer can be readily adapted

to a wide range of plant species and tissue sources. In this chapter

we describe an efﬁ cient, protoplast-based citrus-transformation

system that could be routinely used to transform several important

polyembryonic citrus cultivars that feature robust somatic embryo-

genesis, including important processing sweet oranges and the

popular mandarin cultivar W. Murcott.

The ﬁ rst reports of citrus transformation began to appear more

than two and half decades ago [

150 – 152 ]. Over time, citrus trans-

formation efﬁ ciency has been increased due to continual

improvements in Agrobacterium -mediated methodology and pro-

toplast transformation system, as well as the selection techniques of

the transgenic events. In citrus, the common method of transfor-

mation is Agrobacterium -mediated transformation of stem pieces

(mostly nucellar seedling internodes). This method works best

with seedy polyembryonic cultivars and uses adventitious shoot

induction (organogenesis) as the regeneration pathway. However,

many important citrus cultivars are commercially seedless (zero to

ﬁ ve seeds per fruit) or totally seedless, which makes it difﬁ cult or

impossible to obtain adequate nucellar seedling explants for

Agrobacterium -mediated transformation. Other limitations of

Agrobacterium -mediated citrus transformation include inadequate

susceptibility to Agrobacterium infection and inefﬁ cient plant

regeneration via adventitious shoot-bud induction in certain com-

mercially important cultivars, particularly mandarins. Finally, there

are signiﬁ cant Intellectual Property issues with the use of the com-

mon Agrobacterium -mediated method.

Direct delivery of free DNA molecules into plant protoplasts

has been well documented [

153 ]. Several factors could affect

the efﬁ ciency of free DNA delivery systems, including plasmid

1.3 Citrus

Transformation

Involving Somatic

Embryogenesis

1.3.1 Protoplast

Transformation

Somatic Embryogenesis in Citrus

304

DNA concentration and form, carrier DNA, and treatment and

pretreatment buffers. The delivery of foreign genes into proto-

plasts is usually carried out by electroporation [

154 ] or treatment

with polyethylene glycol ( PEG ) [

130 , 155 ] (Fig. 2 ). The PEG-

mediated transformation is simple and efﬁ cient, allowing a

Fig. 2 GFP selection in protoplast/GFP transformation system. ( a ) protoplasts expressing GFP 24 h after

transformation, ( b ) protoplast derived micro-calli (transformed and non-transformed) under blue light ,

( c ) protoplast derived micro-calli (transformed and non-transformed) under white light , ( d ) transgenic ( green )

and non- transgenic ( red ) somatic embryo s under blue light , ( e ) transgenic ( green ) and non-transgenic ( yel-

low ) somatic embryos under white light , ( f ) transgenic ( green ) and non-transgenic ( red ) somatic embryos

under blue light , ( g ) enlarged transgenic embryo expressing GFP, ( h , i ) transgenic somatic embryo derived

shoots, ( j ) non- transformed shoot, ( k ) micrografting of transgenic shoot onto non-transgenic rootstock, ( l ) GFP

expression in root

Ahmad A. Omar et al.

305

simultaneous processing of many samples, and yields a transformed

cell population with high survival and division rates [

156 ]. The

method utilizes inexpensive supplies and equipment, and helps to

overcome an obstacle of host range limitations of Agrobacterium -

mediated transformation , since DNA uptake by protoplasts is pro-

moted by chemical treatment with PEG. Plant recovery is usually

through the somatic embryo genesis pathway rather than through

organogenesis. Moreover, the transformation method of choice for

plant protoplasts is dependent on a number of factors, including

efﬁ ciency of DNA delivery, toxicity to the cells, ease of use, and

cost and availability of materials. In protoplast transformation sys-

tems, plating and selection methods are important considerations

in the development of stable transgenic plants. The ideal system

should permit easy identiﬁ cation of transformants without the

complications of multiple recovery of single transformation events

or recovery of “false-positives” due to inadequate selection pres-

sure. Therefore using the GFP gene ( green ﬂ uorescent protein ) as

a selectable marker essentially eliminates the problem of multiple

recoveries of single events. Under optimal conditions, up to 50

transformed embryos can be recovered per million input of proto-

plasts (transformation frequency = 0.005 %). The low toxicity,

simplicity, high efﬁ ciency, and low cost of the PEG transformation

method make it an attractive alternative to electroporation as the

method of choice for stable transformation of plant protoplasts.

PEG -mediated gene transfer to citrus protoplasts has proven

to be efﬁ cient, reliable, inexpensive, and a simple method that

works well when using relatively young embryogenic cultures with

good totipotency [

129 , 132 , 157 ]. In this system, large popula-

tions of protoplasts are isolated from embryogenic suspension cul-

tures to increase the likelihood of obtaining an adequate number

of stable independent transformation events. Regeneration of

transgenic plants via somatic embryo genesis is possible under suit-

able in vitro conditions through selection at an early stage of devel-

opment (usually the pro-embryo stage) using GFP gene as a

reporter gene. However, the tissue-culture response may vary

depending on the plant genotype, handling and the condition of

the suspension cells. A major requirement for protoplast transfor-

mation system is the preparation of viable protoplasts. We have

successfully used the procedure described below for gene transfer

to citrus for several cultivars, including “Hamlin” and “Valencia”

sweet oranges, and “W. Murcott” tangor [

9 , 10 , 129 , 132 ]. Cell

suspension s provide an unlimited source of rapidly dividing pro-

toplasts that can be obtained after 12–18 h incubation in enzyme

solution and show a transient expression of introduced genes

within 24 h after transformation. This protocol can be adapted to

a wide range of plant species and tissue sources used for protoplast

preparation.

Somatic Embryogenesis in Citrus

306

Genetic transformation using embryogenic cell suspension cultures

offers a practical alternative to the transformation of epicotyl

explants obtained from germinating seedlings, since almost all

polyembryonic cultivars can be introduced in vitro as embryogenic

cell suspension cultures [

158 ]. Amenability of cell suspension cul-

tures to transformation using Agrobacterium would allow the

transformation of any cultivar that can be introduced as embryo-

genic cell mass es , including specialty seedless sweet oranges or

“Satsuma” mandarins and other difﬁ cult-to-transform cultivars of

the mandarin or lemon group. Our protocol is based on a hygro-

mycin selection regime, as it was observed that kanamycin selec-

tion resulted in erratic and low transgenic embryo production.

Inefﬁ cient kanamycin selection was either due to cells overcoming

the effects of the antibiotic or to the protection of cells from

kanamycin by the surrounding cells [

159 , 160 ]. Successful callus

transformation of sweet oranges and mandarins can be accom-

plished in a selected medium containing 25 mg/L of hygromycin

B. Most material, stocks, and medium are similar to the protoplast

transformation process. Agrobacterium mediated transformation

relies on an active Agrobacterium culture instead of plasmid

DNA as in the protoplast transformation process. Additional mate-

rials required in this protocol are indicated in the protocol section.

A description of the transformation process can also be found in

Dutt and Grosser [

158 ].

2 Materials

1. Fluorescence microscope with FITC ﬁ lters: Zeiss SV11 epiﬂ u-

orescence stereomicroscope equipped with a 100 W mercury

bulb light and a ﬂ uorescein-5-isothiocyanate/ GFP (FITC/

GFP) ﬁ lter set with a 480 nm excitation ﬁ lter and a 515 nm

long-pass emission ﬁ lter (Chroma Technology Corp.,

Brattleboro, VT, USA).

2. Temperature-controlled rotary shaker at 28 ± 2 °C.

3. Laminar ﬂ ow cabinet.

4. pH meter.

5. Autoclave.

6. Sterilized paper plates.

7. Syringe ﬁ lter units, 0.2 μm pore size.

8. Centrifuge with 100–400 × g capability.

9. 40 mL Pyrex tubes.

10. 15 mL Pyrex capped tube.

11. 15-mL round-bottom screw-cap centrifuge tubes.

1.3.2 Agrobacterium -

Mediated Transformation

of Embryogenic Cell

Suspension Cultures

2.1 Equipment

Ahmad A. Omar et al.

307

12. 60 × 15 mm petri dishes.

13. 100 × 20 mm petri dishes.

14. 100 × 15 mm petri dishes.

1. Sterilization solution: 20 % (v/v) commercial bleach solution.

2. BH3 macronutrient stock: 150 g/L KCl, 37 g/L MgSO 4 ·7H

2 O,

15 g/L KH

2 PO

4 , 2 g/L K

2 HPO

4 ; dissolve in H

2 O and store

at 4 °C.

3. Murashige and Tucker (MT) macronutrient stock [

101 ]:

95 g/L KNO

3 , 82.5 g/L NH

4 NO

3 , 18.5 g/L MgSO

4 ·7H

2 O,

7.5 g/L KH

2 PO

4 , 1 g/L K

2 HPO

4 ; dissolve in H

2 O and store

at 4 °C.

4. MT micronutrient stock: 0.62 g/L H

3 BO

3 , 1.68 g/L

MnSO

4 ·H

2 O, 0.86 g/L ZnSO

4 ·7H

2 O, 0.083 g/L KI,

0.025 g/L Na

2 MoO

4 ·2H

2 O, 0.0025 g/L CuSO

4 ·5H

2 O,

0.0025 g/L CoCl

2 ·6H

2 O; dissolve in H

2 O and store at 4 °C.

5. MT vitamin stock: 10 g/L myoinositol, 1 g/L thiamine -HCl,

1 g/L pyridoxine-HCl, 0.5 g/L nicotinic acid , 0.2 g/L gly-

cine ; dissolve in H

2 O and store at 4 °C.

6. MT calcium stock: 29.33 g/L CaCl

2 · 2H

2 O; dissolve in H

2 O

and store at 4 °C.

7. MT iron stock: 7.45 g/L Na

2 EDTA, 5.57 g/L FeSO

4 · 7H

2 O;

dissolve in H

2 O and store at 4 °C.

8. Kinetin (KIN) stock solution: 1 mg/mL; dissolve the powder

in a few drops of 1 N HCl; bring to ﬁ nal volume with H

2 O and

store at 4 °C.

9. BH3 multivitamin stock A: 1 g/L ascorbic acid , 0.5 g/L cal-

cium pantothenate, 0.5 g/L choline chloride, 0.2 g/L folic

acid, 0.1 g/L riboﬂ avin, 0.01 g/L p-aminobenzoic acid,

0.01 g/L biotin; dissolve in H

2 O and store at −20 °C.

10. BH3 multivitamin stock B: 0.01 g/L retinol dissolved in a few

drops of alcohol, 0.01 g/L cholecalciferol dissolved in a few

drops of ethanol, 0.02 g/L vitamin B12; dissolve in H

2 O and

store at −20 °C.

11. BH3 KI stock: 0.83 g/L KI; dissolve in H

2 O and store at 4 °C.

12. BH3 sugar and sugar alcohol stock: 25 g/L fructose, 25 g/L

ribose, 25 g/L xylose, 25 g/L mannose, 25 g/L rhamnose,

25 g/L cellobiose, 25 g/L galactose, 25 g/L mannitol ; dis-

solve in H

2 O and store at −20 °C.

13. BH3 organic acid stock: 2 g/L fumaric acid, 2 g/L citric acid,

2 g/L malic acid, 1 g/L pyruvic acid; dissolve in H

2 O and

store at −20 °C.

2.2 Medium Stock

Solutions

Somatic Embryogenesis in Citrus

308

1. Coumarin (stock solution, 1.46 mg/mL): Dissolve the powder

in warm H

2 O; store at 4 °C.

2. α-Naphthalene acetic acid (NAA; stock solution, 1 mg/10 mL):

Dissolve the powder in a few drops of 5 M NaOH, bring to

ﬁ nal volume with H

2 O and store at 4 °C.

3. 2,4-Dichlorophenoxyacetic acid (2,4-D; stock solution,

1 mg/10 mL): Dissolve the powder in a few drops of 95 %

(v/v) ethanol, bring to ﬁ nal volume with H

2 O; store at 4 °C.

4. 6-Benzylaminopurine (BAP; stock solution, 1 mg/mL):

Dissolve the powder in a few drops of 5 M NaOH, bring to

ﬁ nal volume with H

2 O; store at 4 °C.

5. Gibberellic acid ( GA

3 ; stock solution, 1 mg/mL): Dissolve the

powder in a few drops of 95 % (v/v) ethanol, bring to ﬁ nal

volume with H

2 O, ﬁ lter-sterilize; store in small aliquots at

4 °C; add to the medium after autoclaving and cooling the

medium to 55 °C in a water bath.

The enzyme solution is ﬁ lter sterilized.

1. Calcium chloride (CaCl

2 ·2H

2 O stock solution, 0.98 M):

Dissolve 14.4 g in 100 mL H

2 O and store at −20 °C.

2. Monosodium phosphate (NaH

2 PO

4 stock solution, 37 mM):

Dissolve 0.44 g in 100 mL H

2 O and store at −20 °C.

3. 2 (N-morpholino) ethanesulfonic acid (MES stock solution,

0.246 M): Dissolve 4.8 g in 100 mL H

2 O and store at −20 °C.

4. Enzyme solution: 0.7 M mannitol , 24 mM CaCl

2 , 6.15 mM

MES buffer, 0.92 mM NaH

2 PO

4 , 2 % (w/v) Cellulase

Onozuka RS (Yakult Honsha), 2 % (w/v) Macerozyme R-10

(Yakult Honsha), pH 5.6. To prepare 40 mL of enzyme solu-

tion, dissolve 0.8 g Cellulase Onozuka RS, 0.8 g Macerozyme

R-10 and 5.12 g mannitol in 20 mL H

2 O and add 1 mL of

CaCl

2 · 2H

2 O, NaH

2 PO4 and MES stock solutions; bring vol-

ume to 40 mL with H

2 O, pH to 5.6 using KOH, ﬁ lter-steril-

ize; store at 4 °C for up to 3 weeks.

1. CPW salts stock solution 1: 25 g/L MgSO

4 ·7H

2 O, 10 g/L

KNO

3 , 2.72 g/L KH

2 PO

4 , 0.016 g/L KI, 0.025 ng/L

CuSO

4 ·5H

2 O; dissolve in H

2 O and store at −20 °C.

2. CPW salts stock solution 2: 15 g/L CaCl

2 ·2H

2 O; dissolve in

H

2 O and store at −20 °C.

3. 13 % CPW (13 %, w/v, mannitol solution with CPW salts):

Dissolve 13 g mannitol in 80 mL H

2 O, add 1 mL each of CPW

salts stock solutions 1 and 2; bring volume to 100 mL with

H

2 O, pH to 5.8, ﬁ lter-sterilize; store at room temperature.

2.3 Plant Growth

Regulator Stocks

2.4 Enzyme Stock

Solutions

2.5 CPW Solution

Ahmad A. Omar et al.

309

4. 25 % CPW (25 %, w/v, sucrose solution with CPW salts):

Dissolve 25 g sucrose in 80 mL H

2 O, add 1 mL each of CPW

salts stock solutions 1 and 2; bring to 100 mL with H

2 O, pH

to 5.8, ﬁ lter-sterilize and store at room temperature.

1. PEG 8000 MW (stock solution, 50 %): Place the bottle of PEG

in a water bath at 80 °C until it melts completely, take 250 mL

and mix it with 250 mL H

2 O, add 4 g of resin AG501-X8

(Bio- Rad), stir for 30 min, ﬁ lter out the resin through a layer

of cotton and allow to stand for several hours before use; store

at 4 °C.

2. Polyethylene glycol ( PEG ) working solution: 40 % (w/v) PEG,

0.3 M glucose, 66 mM CaCl

2 ·2H

2 O, pH 6.0. To prepare

100 mL of PEG solution, dissolve 0.97 g CaCl

2 ·2H

2 O and

5.41 g glucose in 10 mL H

2 O, add 80 mL of PEG stock solu-

tion (50 %) and adjust the volume to 100 mL with H

2 O, pH 6;

ﬁ lter-sterilize and store at 4 °C . Check the pH every 2–3

weeks, since this solution acidiﬁ es with time.

3. Elution solutions for PEG removal. Solution A: 0.4 M glucose,

66 mM CaCl

2 ·2H

2 O, 10 % dimethyl sulfoxide ( DMSO ),

pH 6.0. Solution B: 0.3 M glycine adjusted with NaOH pellets

to pH 10.5. Filter-sterilize both solutions; store at room

temperature and mix together (9:1, v:v) immediately prior to

use to avoid precipitation.

1. Any suitable binary vector containing the hygromycin select-

able marker gene for selection in plants. We have had good

success with the pCAMBIA 1300 series of plant transforma-

tion vectors (

www.cambia.org ).

2. Agrobacterium tumefaciens EHA105 stock containing the

appropriate binary vector plasmid (stored in 20 % glycerol

at −80 °C).

3. Solid bacterial growth medium: Yeast Extract Peptone (YEP)

medium (10 g/L peptone, 10 g/L yeast extract, 5 g/L NaCl,

pH 7.0) supplemented with 15 g/L TC agar , 20 mg/L rifam-

picin , and 100 mg/L kanamycin.

4. Liquid bacterial growth medium: YEP medium supplemented

with 20 mg/L rifampicin and 100 mg/L kanamycin.

1. Rifampicin : 20 mg of antibiotic dissolved in 1 mL of DMSO .

2. Acetosyringone: 0.196 mg dissolved in 1 mL of DMSO to

prepare a 100 mM concentration stock solution.

3. Hygromycin sulfate: 50 mg of antibiotic dissolved in 1 mL of

water. The solution sterilized by ﬁ ltration using a 0.2 μm

membrane.

2.6 Protoplast

Transformation

Solutions

2.7 Agrobacterium

Culture Medium

2.8 Suspension Cell

Transformation Stock

Solutions

Somatic Embryogenesis in Citrus

310

4. Timentin and cefotaxime : 400 mg of each antibiotic dissolved

in 1 mL of water. The solution sterilized by ﬁ ltration using a

0.2 μm membrane.

1. EME 0.15 M semisolid medium: 20 mL/L MT macronutrient

stock, 10 mL/L MT micronutrient stock, 10 mL/L MT vita-

min stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 50 g/L sucrose , 0.5 g/L malt extract , 8 g/L agar ,

pH 5.8; autoclave medium and pour into 100 × 20 mm petri

dishes, 35 mL per dish.

2. DOG semisolid medium: Same as EME 0.15 M semisolid

medium plus 5 mg/L kinetin (5 mL kinetin stock solution);

autoclave medium and pour into 100 × 20 mm petri dishes,

35 mL per dish.

3. H+H semisolid medium: 10 mL/L MT macronutrient stock,

5 mL/L BH3 macronutrient stock, 10 mL/L MT micronu-

trient stock, 10 mL/L MT vitamin stock, 15 mL/L MT cal-

cium stock, 5 mL/L MT iron stock, 50 g/L sucrose , 0.5 g/L

malt extract , 1.55 g/L glutamine , 8 g/L agar , pH 5.8; auto-

clave medium and pour into 100 × 20 mm petri dishes, 35 mL

per dish.

1. H+H liquid medium: 10 mL/L MT macronutrient stock,

5 mL/L BH3 macronutrient stock, 10 mL/L MT micronutri-

ent stock, 10 mL/L MT vitamin stock, 15 mL/L MT calcium

stock, 5 mL/L MT iron stock, 35 g/L sucrose , 0.5 g/L malt

extract , 1.55 g/L glutamine , pH 5.8; pour 500 mL aliquots

into 1000 mL glass Erlenmeyer ﬂ asks, autoclave and store at

room temperature.

All protoplast liquid media are ﬁ lter sterilized.

1. BH3 0.6 M liquid medium: 10 mL/L BH3 macronutrient

stock, 10 mL/L MT micronutrient stock, 10 mL/L MT vita-

min stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 2 mL/L BH3 multivitamin stock A, 1 mL/L BH3 mul-

tivitamin stock B, 1 mL/L BH3 KI stock, 10 mL/L BH3

sugar and sugar alcohol stock, 20 mL/L BH3 organic acid

stock, 20 mL/L coconut water , 82 g/L mannitol , 51.3 g/L

sucrose , 3.1 g/L glutamine , 1 g/L malt extract , 0.25 g/L

casein enzyme hydrolysate, pH 5.8; ﬁ lter-sterilize and store at

room temperature.

2.

EME 0.6 M liquid medium: 20 mL/L MT macronutrient

stock, 10 mL/L MT micronutrient stock, 10 mL/L MT vita-

min stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 205.4 g/L sucrose , 0.5 g/L malt extract

, pH 5.8; ﬁ lter-

sterilize and store at room temperature.

2.9 Callus-

Induction Media

2.10 Cell Suspension

Maintenance Medium

2.11 Protoplast

Isolation and Culture

Media

Ahmad A. Omar et al.

311

1. EME 0.15 M liquid medium: 20 mL/L MT macronutrient

stock, 10 mL/L MT micronutrient stock, 10 mL/L MT vita-

min stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 50 g/L sucrose , 0.5 g/L malt extract , pH 5.8; ﬁ lter-

sterilize and store at room temperature.

2. EME–malt 0.15 M liquid medium: 20 mL/L MT macronutri-

ent stock, 10 mL/L MT micronutrient stock, 10 mL/L MT

vitamin stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 50 g/L maltose , 0.5 g/L malt extract , pH 5.8; ﬁ lter-

sterilize and store at room temperature.

3. EME–malt 0.15 M semisolid medium: 20 mL/L MT macro-

nutrient stock, 10 mL/L MT micronutrient stock, 10 mL/L

MT vitamin stock, 15 mL/L MT calcium stock, 5 mL/L MT

iron stock, 50 g/L maltose , 0.5 g/L malt extract , 8 g/L agar ,

pH 5.8; autoclave medium and pour into 100 × 20 mm petri

dishes, 35 mL per dish.

4. EME 1500 semisolid medium: 20 mL/L MT macronutrient

stock, 10 mL/L MT micronutrient stock, 10 mL/L MT

vitamin stock, 15 mL/L MT calcium stock, 5 mL/L MT iron

stock, 50 g/L sucrose , 1.5 g/L malt extract , 8 g/L agar ,

pH 5.8; autoclave medium and pour into 100 × 20 mm petri

dishes, 35 mL per dish.

5. B+ semisolid medium: 20 mL/L MT macronutrient stock,

10 mL/L MT micronutrient stock, 10 mL/L MT vitamin

stock, 15 mL/L MT calcium stock, 5 mL/L MT iron stock,

25 g/L sucrose , 20 mL/L coconut water , 14.6 mg/L couma-

rin (10 mL coumarin stock), 0.02 mg/L NAA (200 μl NAA

stock), 1 mg/L GA

3 (add 1 mL GA

3 stock after medium is

autoclaved and cooled to 55 °C in water bath), 8 g/L agar ,

pH 5.8; autoclave medium and pour into 100 × 20 mm petri

dishes, 35 mL per dish.

6. DBA3 semisolid medium: 20 mL/L MT macronutrient stock,

10 mL/L MT micronutrient stock, 10 mL/L MT vitamin

stock, 15 mL/L MT calcium stock, 5 mL/L MT iron stock,

25 g/L sucrose , 1.5 g/L malt extract , 20 mL/L coconut

water , 0.01 mg/L 2,4-D (100 μl 2,4-D stock), 3 mg/L BAP

(3 mL BAP stock); 8 g/L agar , pH 5.8; autoclave medium and

pour into 100 × 20 mm petri dishes, 35 mL per dish.

7. RMAN medium (Root induction and propagation): 10 mL/L

MT macronutrient stock, 5 mL/L MT micronutrient stock,

5 mL/L MT vitamin stock, 15 mL/L MT calcium stock,

5 mL/L MT iron stock, 25 g/L sucrose , 0.5 g/L activated

charcoal , 8 g/L agar , 0.02 mg/L NAA (200 μl NAA stock

solution), pH 5.8; autoclave medium and pour into sterile

Magenta GA-7 boxes, 80 mL per box.

2.12 Protoplast

Culture and Plant

Regeneration Media

Somatic Embryogenesis in Citrus

312

1. EME– sucrose 0.15 M semisolid medium supplemented with

Acetosyringone: 20 mL/L MT macronutrient stock, 10 mL/L

MT micronutrient stock, 10 mL/L MT vitamin stock,

15 mL/L MT calcium stock, 5 mL/L MT iron stock, 50 g/L

sucrose, 0.5 g/L malt extract , 8 g/L agar , pH 5.8; autoclave

medium, add 1 mL/L acetosyringone stock solution to par-

tially cooled medium and pour into 100 × 20 mm petri dishes,

35 mL per dish.

2. EME– sucrose 0.15 M liquid medium: 20 mL/L MT macro-

nutrient stock, 10 mL/L MT micronutrient stock, 10 mL/L

MT vitamin stock, 15 mL/L MT calcium stock, 5 mL/L MT

iron stock, 50 g/L sucrose, 0.5 g/L malt extract . Pour into

250 mL bottles before autoclaving.

3. EME– maltose 0.15 M semisolid medium supplemented with

antibiotics: 20 mL/L MT macronutrient stock, 10 mL/L MT

micronutrient stock, 10 mL/L MT vitamin stock, 15 mL/L

MT calcium stock, 5 mL/L MT iron stock, 50 g/L maltose,

0.5 g/L malt extract , 8 g/L agar , pH 5.8; autoclave medium,

add 1 mL/L timentin, 1 mL/L cefotaxime and 500 mg/L

hygromycin stock solutions to partially cooled medium, and

pour into 100 × 20 mm petri dishes, 35 mL per dish.

4. EME 1500 semisolid medium supplemented with antibiotics:

20 mL/L MT macronutrient stock, 10 mL/L MT micronutri-

ent stock, 10 mL/L MT vitamin stock, 15 mL/L MT calcium

stock, 5 mL/L MT iron stock, 50 g/L sucrose , 1.5 g/L malt

extract , 8 g/L agar , pH 5.8; autoclave medium, add 0.5 mL/L

timentin, 0.5 mL/L cefotaxime and 500 mg/L hygromycin

stock solutions to partially cooled medium, and pour into

100 × 20 mm Petri dishes, 35 mL per dish.

5. B+ semisolid medium supplemented with antibiotics: 20 mL/L

MT macronutrient stock, 10 mL/L MT micronutrient stock,

10 mL/L MT vitamin stock, 15 mL/L MT calcium stock,

5 mL/L MT iron stock, 25 g/L sucrose , 20 mL/L coconut

water , 14.6 mg/L coumarin (10 mL coumarin stock),

0.02 mg/L NAA (200 μl NAA stock), 1 mg/L GA

3 (add 1 mL

GA

3 stock solution after medium is autoclaved and cooled to

55 °C in water bath), 8 g/L agar , pH 5.8; autoclave medium,

add 0.5 mL/L timentin stock solution to partially cooled

medium and pour into 100 × 20 mm petri dishes, 35 mL per

dish.

6. DBA3 semisolid medium supplemented with antibiotics:

20 mL/L MT macronutrient stock, 10 mL/L MT micronutri-

ent stock, 10 mL/L MT vitamin stock, 15 mL/L MT calcium

stock, 5 mL/L MT iron stock, 25 g/L sucrose , 1.5 g/L malt

extract , 20 mL/L coconut water , 0.01 mg/L 2,4-D (100 μl

2,4-D stock solution), 3 mg/L BAP (3 mL BAP stock solu-

2.13 Suspension

Culture

Transformation

and Plant

Regeneration Media

Ahmad A. Omar et al.

313

tion); 8 g/L agar , pH 5.8; autoclave medium, add 0.5 mL/L

timentin stock solution to partially cooled medium and pour

into 100 × 20 mm petri dishes, 35 mL per dish.

7. RMAN medium supplemented with antibiotics: 10 mL/L MT

macronutrient stock, 5 mL/L MT micronutrient stock,

5 mL/L MT vitamin stock, 15 mL/L MT calcium stock,

5 mL/L MT iron stock, 25 g/L sucrose , 0.5 g/L activated

charcoal , 8 g/L agar , 0.02 mg/L NAA (200 μl NAA stock

solution), pH 5.8; autoclave medium, add 0.5 mL/L timentin

stock solution to partially cooled medium and pour into sterile

Magenta GA-7 boxes, 80 mL per box.

3 Methods

1. Immerse harvested immature fruit in sterilization solution in a

beaker for 30 min.

2. Using sterile tongs, place fruit on sterilized paper plates in a

laminar ﬂ ow hood.

3. Using a sterile surgical blade, make an equatorial cut, 1–2 cm

deep, and open the fruit.

4. With sterile forceps, extract ovules and place them onto callus-

induction medium (EME 0.15 M, H+H or DOG).

5. Incubate extracted ovules in the dark at 28 ± 2 °C and transfer

them every 3–4 weeks to new callus-induction medium until

embryogenic (yellow and friable) callus emerges from the

ovules.

6. To maintain long-term cultures, transfer embryogenic undif-

ferentiated calli ( see Note 1 ) onto new medium every 4–6

weeks and incubate under the same conditions.

7. To initiate cell suspension s from embryogenic undifferentiated

nucellus -derived callus, take approx. 2 g of calli from callus-

induction medium and transfer to 125 mL Erlenmeyer ﬂ asks,

each containing 20 mL of H+H liquid medium.

8. Shake the cell suspension cultures on a rotary shaker at 125 rpm

under a 16 h photoperiod (70 μmol m

−2 s

−1 ) at 28 ± 2 °C.

9. After 1 week, add 10 mL of new H+H liquid medium to

Erlenmeyer ﬂ asks and return back to the shaker.

10. After one more week, add 20 mL of new H+H liquid medium

to Erlenmeyer ﬂ asks and return back to the shaker.

11. Subculture established embryogenic cell suspension cultures

every 2 weeks by removing 20 mL from the culture and replac-

ing with 20 mL fresh aliquots of H+H liquid medium; shake at

125 rpm and incubate under the same conditions.

3.1 Protoplast

Transformation [ 9 ,

129 ]: Initiation

and Maintenance

of Embryogenic

(Callus and Cell

Suspension) Cultures

Somatic Embryogenesis in Citrus

314

1. Transfer 1–2 g of friable callus into a 60 × 15 mm petri dish. If

using a suspension as a source for embryogenic cells (s ee Note 2 )

transfer approx. 2 mL of suspension ( see Note 3 ) with a wide-

mouth pipette.

2. Drain off the liquid medium using a Pasteur pipette.

3. Resuspend the cells in a mixture of 2.5 mL 0.6 M BH3 liquid

medium and 1.5 mL enzyme solution ( see Note 4 ).

4. Seal petri dishes with Paraﬁ lm and incubate overnight (15–20 h)

at 28 °C on a rotary shaker at 25–30 rpm in the dark.

1. Following overnight incubation, pass enzymatic preparations

through a sterile 45 μm nylon mesh sieve ( see Note 5 ) to

remove undigested tissues and other cellular debris; collect the

ﬁ ltrate in 40 mL Pyrex tubes.

2. Transfer the protoplast-containing ﬁ ltrate ( see Note 6 ) to a

15 mL calibrated screw-cap centrifuge tube.

3. Centrifuge at 900 rpm for 10 min.

4. Remove the supernatant with a Pasteur pipette and gently

resuspend the protoplast pellet in 5 mL of 25 % CPW

solution.

5. Slowly pipette 2 mL of 13 % CPW solution directly on top of

the sucrose layer. Avoid mixing the two layers.

6. Centrifuge at 900 rpm for 10 min.

7. Only viable protoplasts ( see Note 7 ) form a band at the inter-

face between the sucrose and the mannitol layers.

8. Remove the protoplasts ( see Note 8 ) from this interface with a

Pasteur pipette and resuspend them in 10 mL of BH3 0.6 M

liquid medium (using a new screw-cap centrifuge tube).

9. Centrifuge at 900 rpm for 10 min.

10. Remove the supernatant and gently resuspend the pellet in

10 mL of BH3 0.6 M medium ( see Note 9 ).

11. Centrifuge at 900 rpm for 10 min.

12. Remove the supernatant and gently resuspend the pellet in

10 mL of BH3 0.6 M medium.

13. Centrifuge at 900 rpm for 10 min.

14. Remove the supernatant and resuspend the pellet into 5 mL

BH3 0.6 M.

15. Determine protoplast density using a hemocytometer

( see Note 10 ).

16. Centrifuge at 900 rpm for 10 min.

17. Remove the supernatant and resuspend the pellet into BH3

0.6 M to reach 4 × 10

6 protoplasts/mL.

3.2 Protoplast

Transformation:

Preparation

and Enzymatic

Incubation of Cultures

from Embryogenic

Callus

3.3 Protoplast

Transformation:

Protoplast Isolation

and Puriﬁ cation

[

9 , 129 ]

Ahmad A. Omar et al.

315

1. In a 15 mL round-bottom screw- cap centrifuge tubes

( see Note 12 ) add 0.5 mL of protoplast suspension (2 × 10

6

protoplasts/mL).

2. Add 30–40 μg plasmid DNA ( see Note 13 ) and gently mix

well by gentle agitation.

3. Immediately add 0.5 mL of PEG solution directly into the

center of the tube to give the desired ﬁ nal PEG concentration

(20 %) ( see Note 14 ), allowing the PEG to mix with the pro-

toplast suspension by gentle agitation ( see Note 15 ).

4. After 25–30 min, add 0.5 mL of A + B solution (9:1, v:v) into

each transformation tube, but this time gently and slowly onto

the inside edge of the tube, trying not to agitate the fragile

transforming protoplasts.

5. After another incubation period of 25–30 min, gently add

1 mL of BH3 0.6 M medium onto the inside edge of the tube,

again trying not to disturb the protoplasts.

6. After incubating for an additional 10 min, dilute the protoplast

suspension with four 1-mL aliquots of BH3 0.6 M at 5 min

intervals onto the inside edge of the tube, again trying not to

disturb the protoplasts.

7. Cap and seal the tube with Paraﬁ lm.

8. Centrifuge at 700 rpm for 5 min.

9. Carefully, remove supernatant, add 2 mL BH3 0.6 M medium

and gently resuspend the protoplast.

10. Centrifuge at 700 rpm for 5 min.

11. Carefully, remove supernatant, add 2 mL BH3 0.6 M medium

and gently resuspend the protoplast ( see Note 16 ).

12. Repeat steps 10

and 11 one more time, carefully avoiding the

loss of protoplasts.

13. Finally, add 1–1.5 mL of a 1:1 (v:v) mixture of BH3 0.6 M and

EME 0.6 M liquid media to each tube, gently resuspend the

protoplast.

14. Transfer the suspended protoplast into 60 × 15 mm petri dishes

and spread into a thin layer by gently swirling the petri dishes

( see Note 17 ).

15. Seal the dishes with Paraﬁ lm and culture in the dark at 28 ± 2 °C

for 4–6 weeks ( see Note 18 ).

16. Check GFP expression 48 h after transformation ( see Note 19 )

using Zeiss SV11 epiﬂ uorescence stereomicroscope and return

the dishes back in the dark at 28 ± 2 °C (Fig.

2 ).

3.4 Protoplast

Transformation:

Polyethylene Glycol

( PEG )-Induced

Protoplast

Transformation [

129 ]

( see Note 11 )

Somatic Embryogenesis in Citrus

316

1. After 4–6 weeks of incubation ( see Note 20 ), supplement

cultures of transformed protoplasts with new medium con-

taining reduced osmoticum. Accomplish this by adding 10–12

drops of 1:1:1 (by volume) mixture of BH3 0.6 M, EME

0.6 M, and EME 0.15 M liquid media.

2. Incubate cultures for another 2 weeks in low light

(20 μmol m

−2 s

−1 intensity) with a 16 h photoperiod at 28 ± 2 °C.

3. Accomplish another reduction of osmoticum in the cultures by

the following steps: add 2 mL of 1:2 (v:v) mixture of BH3

0.6 M and EME-malt 0.15 M liquid media to each dish of

transformed-treated protoplasts.

4. Immediately pour the entire contents onto petri dishes with

agar -solidiﬁ ed EME-malt 0.15 M medium and swirl gently

each dish in order to spread the liquid containing protoplast-

derived colonies evenly over the entire semisolid agar surface.

5. Incubate cultures with a 16 h photoperiod (70 μmol m

−2 s

−1

intensity) at 28 ± 2 °C and, from this point until somatic

hybrid s are planted in compost, keep the cultures under the

same growth conditions.

6. Transfer regenerated somatic embryo s as soon as they appear

from callus colonies to new agar -solidiﬁ ed EME-malt 0.15 M

medium ( see Note 21 ).

7. After 3–4 weeks, move small somatic embryo s to semis solid

EME 1500 medium for enlargement and germination .

8. Move the germinated embryos to semisolid B+ medium for

axis elongation.

9. Dissect abnormal embryos that fail to germinate into large sec-

tions and place on DBA3 medium for shoot induction.

10. Transfer all resulting GFP positive shoots into RMAN medium

to induce rooting ( see Note 22 ) (Fig.

2 ).

11. Transfer rooted plants into peat based potting mixture in the

greenhouse and cover with rigid clear plastic for 3–4 weeks

maintaining high humidity.

12. Remove the plastic covers following this period of

acclimatization.

13. After having an established plant with 3–4 leaves start molecu-

lar analysis ( see Note 23 ).

1. Obtain Agrobacterium cultures kept in a −80 °C freezer and

thaw.

2. Remove a loopful of bacteria from each thawed culture, and

streak it on an individual YEP plate.

3. Incubate plates at 28 °C for 2 days.

4. Use a single bacterial colony and inoculate a ﬂ ask of 25 mL

liquid YEP medium containing appropriate antibiotics.

3.5 Protoplast

Transformation:

Protoplast Culture

and Plant

Regeneration

3.6 Suspension Cell

Culture

Transformation [

158 ]:

Agrobacterium

Preparation

and Culture

Transformation

Ahmad A. Omar et al.

317

5. Culture for 24 h at 28 °C.

6. Transfer a 3–5 mL overnight aliquot into fresh 25 mL liquid

YEP medium containing appropriate antibiotics.

7. Culture for 3–4 h at 26 °C.

8. Centrifuge cells at 6000 rpm for 8 min at 25 °C.

9. Resuspend cells in 25 mL liquid EME- sucrose medium.

10. Prior to use in transformation, measure the optical density

(OD) of cultures and adjust to 0.3.

11. Transfer 20 mL of cell suspension cultures into a 100 × 15 mm

petri dish. Drain off the liquid medium using a Pasteur pipette.

12. Transfer bacterial solution into the suspension cells for 20 min

with frequent and gentle agitation.

13. Blot cell suspension cultures on sterile paper towels and trans-

fer onto semisolid EME- sucrose medium supplemented with

acetosyringone.

14. Incubate in the dark at 25 °C for 5 days.

1. Transfer putative transgenic cells onto EME + maltose

embryo production medium supplemented with appropri-

ate antibiotics.

2. Maintain cultures either in the dark or under low light

(20 μmol m

−2 s

−1 intensity) condition.

3. After 4–6 weeks in this medium, transfer cells into fresh

medium. At this stage add 2 mL of 1:2 (v:v) mixture of BH3

0.6 M and EME-malt 0.15 M liquid media to each dish of

transformed-suspension cells. Supplement the 1:2 mixture

with 200 mg/L timentin and 25 mg/L hygromycin.

4. Transfer regenerated somatic embryo s as soon as they appear

from callus colonies to new agar -solidiﬁ ed antibiotic supple-

mented EME- maltose medium.

5. After 3–4 weeks, move small somatic embryo s to semi solid

EME 1500 antibiotic supplemented medium for enlargement

and germination .

6. Move the germinated embryos to semisolid antibiotic supple-

mented B+ medium for axis elongation.

7. Dissect abnormal embryos that fail to germinate into large sec-

tions and place on antibiotic supplemented DBA3 medium for

shoot induction.

8. Transfer all resulting shoots into RMAN medium to induce

rooting.

9. Transfer rooted plants into a peat based potting mixture in the

greenhouse and cover with rigid clear plastic for 3–4 weeks

maintaining high humidity.

3.7 Suspension Cell

Culture

Transformation:

Selection of Putative

Transformed Embryos

and Regeneration

of Transformed Plants

Somatic Embryogenesis in Citrus

318

10. Remove the plastic covers following this period of

acclimatization.

11. Established plants with 3–4 leaves can then be subjected to

appropriate molecular analysis to determine gene insertion.

4 Notes

1. Since the nucellar callus has high embryogenic capacity, the

best way to maintain the long-term callus in an undifferenti-

ated state is to visually select and subculture only white/yellow

friable callus. Differentiated callus types and organized tissues

should be discarded.

2. Cultured embryogenic cells used for protoplast isolation

should be in the log phase of growth. For consistent results,

maintain uniform growth conditions for the cell suspension ,

because the physiological state of the suspension cells is an

important factor inﬂ uencing protoplast yield, quality and

transformation efﬁ ciency. Use 5–12 day-old suspensions from

a 2 week subculture cycle, or 7–21 day-old callus from a 4

week subculture cycle.

3. Cell suspension morphology differs from one genotype to

another, thus we recommend using a volume of suspension

that approximates 1 g fresh weight of callus.

4. Best release of protoplasts is obtained with freshly prepared

digestion enzymes, do not store enzyme solution more than

2 weeks.

5. Nylon mesh is sealed to a 4 cm long plastic cylindrical tube

made from a plastic syringe. In order to make a similar piece of

equipment, take a 30 mL plastic syringe, cut it at the 25 mL

mark and keep the upper part with wings. Place a nylon mem-

brane on a preheated hot plate beneath the cylindrical tube

and seal the two parts.

6. Protoplasts are fragile, thus take extra care when ﬁ ltering the

protoplast/enzyme solution and later when centrifuging and

resuspending protoplasts. When being transferred from one

tube to another it is important that the protoplasts are drawn

gently into the Pasteur pipette and dispensed slowly down the

inside wall of the receiving centrifuge tube. Also, when resus-

pending pellets of protoplasts with different solutions, ensure

a gentle technique of breaking clumps by introducing small

bubbles of air with a Pasteur pipette, instead of sucking sus-

pensions in and out of the pipette. Mishandling of the proto-

plasts can affect their integrity and thereby affect the efﬁ ciency

of the procedure.

7. If, after isolation and puriﬁ cation, a good yield of protoplasts

(5–10 × 10 6 protoplasts/incubation plate) is not obtained, it

Ahmad A. Omar et al.

319

may be necessary to vary both the enzyme concentration and

length of incubation time to optimize digestion efﬁ ciency.

8. When recovering protoplasts from the sucrose - mannitol gradi-

ent take as little of the sucrose as possible with the protoplasts.

Retention of too much sucrose makes it difﬁ cult to pellet the

protoplasts at later steps.

9. The washing step that removes the enzymes seems to have a

greater bearing on the transformation efﬁ ciency, because pro-

toplast samples that have not been washed very well always

yield lower transformation, division, and survival rates. It is

recommended to repeat steps 10 and 11 in method

3.3 until

a tight clean pellet is obtained.

10. Determine the protoplast density using a hemocytometer. If

the number of protoplasts exceeds 100 cells/square in the

hemocytometer, dilute the protoplast suspension to obtain

accurate counting.

11. Perform protoplast transformation ( PEG -induced method)

within 1–2 h (preferably immediately) after protoplast isola-

tion, since protoplasts start to regenerate cell wall s as soon as

they are rinsed from the enzyme solution. Cell wall regenera-

tion may hinder transformation.

12. The number of tubes is determined by the total volume of

mixed protoplasts at 4 × 10

6 protoplasts/mL.

13. The DNA should be sterile (ethanol-precipitated and dissolved

in sterile water). Do not incubate DNA for too long with the

protoplasts because it may result in lower transformation efﬁ -

ciency due to nuclease digestion. The DNA concentration

should be at least 1 μg/μl, to minimize the added volume.

In the co-transformation you will add two DNA plasmids one

for the gene of interest and the other for the reporter gene.

In the direct transformation you will add one DNA plasmid

which contain both the gene of interest and the reporter gene

in one construct.

14. Using high PEG concentration could reduce the transforma-

tion frequency due to either PEG toxicity, lower ﬁ nal DNA

concentration, or a combination of these two factors. Use only

freshly prepared and ﬁ lter-sterilized PEG solution (do not

autoclave). Check the pH periodically. The PEG should be

added immediately after DNA addition as protoplasts are a rich

source of nucleases (secretion and release by breakage) that

may hydrolyze the DNA.

15. The protoplast/ PEG solution may be agitated gently every

5 min for 30 min. A certain proportion of protoplasts will

invariably break during and following the PEG treatment. The

debris of dead cells is detrimental for a continued liquid culture

of surviving protoplasts, thus try to handle the protoplast/

PEG culture very gently to reduce this phenomena.

Somatic Embryogenesis in Citrus

320

16. Washing the protoplasts three times is very important to

remove all the PEG and A:B solution.

17. In each transforming dish, protoplasts are plated at a density of

approx. 1–1.5 × 10

6 protoplasts/mL of culture medium. In

order to retain viability and induce cell division, transformed

protoplasts have to be plated in thin-layer culture at high cell

density. In the case of citrus protoplasts, the best results are

obtained when the cell density exceeds 1 × 10

6 protoplasts/mL

of medium. If necessary, determine and adjust protoplast den-

sity using a hemocytometer.

18. Check the cultures every week to evaluate the rate of colony

development. In the event of fast colony development feeding

can begin as early as 10–14 days after transformation as fol-

lows: add 6–8 drops of liquid 1:1:1 of BH3 0.6 M–EME

0.6 M–EME 0.146 M medium to reduce the osmotic pressure

and incubate the cultures again in the dark at 28 °C without

agitation. Ten to 14 days later add another 6–8 drops of liquid

1:1:1 of BH3 0.6 M–EME 0.6 M–EME 0.146 M medium.

Depending on the quality of protoplast preparations, up to

75 % of the protoplasts survive and 20–40 % of cells will

undergo divisions during the ﬁ rst 7–10 days of culture. After

14 days of culture, the dividing cells should form colonies of

2–16 cells.

19. Optimal ﬂ uorescence is observable only after 48 h, although

some transformed protoplasts start exhibiting GFP ﬂ uores-

cence 24 h after transformation.

20. In the case of slow developing colonies, begin feeding 4–6

weeks after transformation.

21. Take care of those GFP positive embryos developing faster and

separate them from the rest. Transfer them earlier for regenera-

tion to plants. Faster-developing embryos can rapidly produce

healthy (normal) plants. It is recommended to place a few

(6–8) embryos together onto cellulose acetate ﬁ lter paper for

rapid and normal development.

22. In certain leaf pieces, the green ﬂ uorescence can be entirely

masked by the chlorophyll pigment. In case of doubts about

the transgenic nature of regenerated plants, a small portion of

leaves may be used to make protoplasts in 1 mL of enzyme

solution and the protoplasts may directly be observed as they

are released. To accelerate the propagation of the transgenic

shoots, you can use any available grafting technique, either

shoot tip grafting onto a greenhouse growing rootstock or

in vitro micro-grafting on seedling rootstock [

129 ].

23. Molecular analysis (PCR, Southern and Western analysis)

should conﬁ rm the integration and expression of the transgene

in the citrus genome.

Ahmad A. Omar et al.

321

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_14, © Springer Science+Business Media New York 2016

Chapter 14

Somatic Embryogenesis Induction and Plant Regeneration

in Strawberry Tree ( Arbutus unedo L.)

João F. Martins , Sandra I. Correia , and Jorge M. Canhoto

Abstract

Somatic embryogenesis is a powerful tool both for cloning and studies of genetic transformation and

embryo development. Most protocols for somatic embryogenesis induction start from zygotic embryos or

embryonic-derived tissues which do not allow the propagation of elite trees. In the present study, a reliable

protocol for somatic embryogenesis induction from adult trees of strawberry tree is described. Leaves from

in vitro proliferating shoots were used to induce somatic embryo formation on a medium containing an

auxin and a cytokinin. Somatic embryos germinated in a plant growth regulator-free medium.

Key words Epicormic shoot s , Ericaceae , Fruit , Germination , In vitro , Neglected crops , Shoot prolif-

eration , Somatic embryo

1 Introduction

Arbutus unedo L. is an Ericaceae species commonly known as

strawberry tree . It grows on acidic, rocky, and well-drained soils

[

1 , 2 ] and can withstand low (until −12 °C) temperatures [ 1 ] as

well as dry conditions [

1 ]. A. unedo individuals grow spontane-

ously (Fig.

1a ) in several countries of the Mediterranean Basin,

from Spain to Turkey, as well as in North Africa, Mediterranean

Islands, and Atlantic Coast, including Ireland and Portugal [

1 , 2 ].

Strawberry tree is a small perennial shrub or tree (Fig.

1b ) that

usually grows up to 3 m [

3 ] with a spreading habit and gray-brown

bark. The edible fruit (Fig.

1c ) is a spherical berry, with about 2 cm

diameter, covered with conical papillae and enclosing 10–50 small

seed whereas the ﬂ owers are hermaphrodite, bell shaped, whitish

to slightly pink, and organized in hanging panicles (Fig.

1d ), [ 4 ].

The reproductive cycle is long with fruits taking a year to ripe, and,

during several months of the year, both ﬂ owers and fruits are pres-

ent in the same tree, making the species a very attractive ornamen-

tal plant. From an ecological perspective, strawberry tree is a very

important species in Mediterranean ecosystems avoiding erosion,

330

Fig. 1 Aspects of strawberry tree. ( a ) Field-growing trees in the North of Portugal, on the slopes of a hill near a

dam (Google Earth information: 41°45′55″N, 8°11′89″, altitude 639 m). Adult tree ( b ), the fruits ( c ), and the

ﬂ owers ( d ). Fruits in fermentation for the production of the spirit “medronheira” ( e )

providing food for fauna and helping recover marginal lands [ 1 , 4 ].

Its ability to regenerate after ﬁ res is a feature that makes the species

interesting for reforestation programs, especially in southern

Europe countries such as Portugal, Spain, Italy, and Greece, where

forest ﬁ res are common [

5 ]. Once considered a “Neglected or

Underutilized Crop” (

www.cropsforthefuture.org ), the impor-

João F. Martins et al.

331

tance of strawberry tree is growing [ 3 , 4 ]. In a context where some

of the most important forest species in southern Europe, such as

pine and eucalyptus, are suffering from several diseases, the demand

for strawberry tree by producers and stakeholders is increasing,

particularly in Portugal [

3 , 4 ]. The fruits are commonly used in the

manufacture of traditional products such as jam and jelly [

5 ].

However, its main application is for the production of an alcoholic

distillate [

6 ], known in Portugal as medronheira (Fig. 1e ).

The propagation of strawberry tree can be achieved through

the use of conventional methods of vegetative propagation such as

cuttings [

7 ] or by seeds [ 8 , 9 ]. Seeds do not assure true-to-type

propagation and particular characteristics can be lost. Assays of

vegetative propagation can be made by conventional vegetative

propagation methods such as rooting or grafting. However, the

frequencies of rooting are quite low, especially when mature cut-

tings are used [

10 , 11 ], and elite genotypes are unavailable for

grafting. In vitro tissue culture techniques have been applied to the

propagation of strawberry tree, in particular axillary shoot prolif-

eration [

11 , 12 ]. However, somatic embryo genesis has much more

potential for cloning than other micropropagation techniques

since somatic embryos are easier to handle than other propagules

and can be obtained in large amounts from a single explant [

13 ,

14 ]. Somatic embryo genesis has great potential for genetic trans-

formation and cryopreservation of desirable selected lines [

15 ,

16 ]. Moreover, somatic embryo induction and development serve

as a model to understand the physiological and genetic factors con-

trolling the different steps of embryo development [

17 , 18 ].

Previous works on A. unedo have shown that somatic embryogen-

esis can be achieved from adult material [

19 , 20 ].

Here, a protocol for somatic embryo genesis induction and

plant regeneration from A. unedo trees is described. To overcome

the lack of potential of adult tissues for somatic embryogenesis,

an indirect approach was attempted in which shoots from selected

adult trees were ﬁ rst established in vitro through axillary shoot

proliferation and then somatic embryogenesis was induced in

leaves from these shoots. This protocol of somatic embryogenesis

induction and plant regeneration can also be applied to the prop-

agation of Arbutus canariensis Duham, a species quite similar to

A. unedo .

2 Materials

Somatic embryo genesis in A. unedo can be induced from leaves of

in vitro propagated shoots, established from adult or juvenile (i.e.,

not yet in the reproductive phase) plants or from seedlings. The

methodology is the same but shoots from adult trees are of known

genotypes allowing the propagation of selected trees. Hence, the

Somatic Embryogenesis in Strawberry Tree

332

following methodology is based on experiments with leaves from

in vitro developing shoots established from adult plants.

1. Selected adult trees (Fig.

1a ).

2. Semi-woody branches ( see Note 1 ), collected from the selected

plants (30–40 cm length).

3. Epicormic shoot s (Fig.

2a ) of 2–4 cm, collected from these

branches.

4. Proliferating shoots (Fig.

2b ).

5. Leaves from proliferating shoots (Fig.

2c ).

1. Semisolid medium for culture establishment (SP medium):

Major salts from the Anderson medium [

21 ], micronutrients

from the Murashige and Skoog [

22 ] medium (without KI),

and organic compounds of the De Fossard medium [

23 ]. Add

0.087 M sucrose and 8.8 μM BA (6-benzyladenine, Sigma

Chemical Company, St. Louis, MO, USA).

2. Induction medium (IM): This medium contains the same

components of the SP medium and the following growth reg-

ulators: 8.8 μM BA and 26.8 μM NAA (1-naphthaleneacetic

acid, Sigma Chemical Company, St. Louis, MO, USA).

3. Somatic embryo germination medium (GM): Knop [

24 ] major

salts, micronutrients of the Murashige and Skoog medium [

22 ],

vitamins (without riboﬂ avin) of the De Fossard medium [

23 ],

0.044 M sucrose and 1 % (w/v) activated charcoal ( Merck

KGaA, Darmstadt, Germany).

4. Add 0.6 % (w/v) agar (Panreac, Spain, or equivalent) to all

media before autoclaving at 121 °C for 20 min (800–1100 g/

cm gel strength after autoclaving). Adjust the pH of all media

to 5.7 using KOH or HCl diluted solutions (0.01–1 M) before

autoclaving and agar addition ( see Note 2 ).

3 Methods

1. Remove the leaves from semi-woody branch segments and

wash them in running water to remove major detritus. Spray

washed branches with a fungicide solution (Benlate or equiva-

lent, 6 % w/v), and set upright in jars containing water to allow

the development of axillary shoots in a growth cabinet at 20 °C

and 80–90 % relative humidity, under a 16 h daily illumination

regime of 15–20 μmol/m

2 /s photosynthetically active radia-

tion (PAR, cool-white ﬂ uorescent lamps) ( see Note 3 ). Change

the water every 2 days to avoid fungi growth ( see Note 4 ).

2.1 Plant Material

2.2 Culture Media

João F. Martins et al.

333

Fig. 2 Formation of epicormic shoot s and shoot proliferation. ( a ) Epicormic shoot s

(ep) developing from a branch after 1.5 month. ( b ) Shoot proliferation on medium SP.

( c ) Wounded ( arrows ) leaves at the time of the culture for somatic embryo genesis

induction

Somatic Embryogenesis in Strawberry Tree

334

2. Isolate the shoot apex (0.5 cm) and nodal segments ( see Note 5 )

from the epicormic shoot s (3–4 cm length). Remove the leaves

and wash with detergent (2–3 drops of Tween 20) in a volume

of 100 mL of water.

3. Transfer the explants to an ethanol ( see Note 6 ) solution (70 %

v/v) during 30 s.

4. Wash three times with sterilized water, and transfer to a 5 %

(w/v) calcium hypochlorite solution containing 2–3 drops of

Tween 20 under stirring, during 15 min ( see Note 7 ).

5. Wash three times with sterilized water to remove the excess of

hypochlorite.

6. In a laminar ﬂ ow chamber, transfer shoot apices and nodal seg-

ments of epicormic shoot s to test tubes (15 × 2.2 cm) containing

15 mL/tube of SP medium ( see Note 8 ). Place one apex or nodal

segment per test tube for culture establishment ( see Note 9 ).

7. Keep the cultures in a growth chamber under a 16 h photope-

riod of 15–20 μmol/m

2 /s (cool-white ﬂ uorescent lamps) at

25 °C ( see Note 10 ).

8. Following establishment, shoots can be subcultured to obtain

large amounts of leaves for further assays. Subcultures must be

carried in the same conditions than in vitro establishment.

9. Leaves (0.4–0.8 cm length) from proliferating shoots (3–4 cm)

are used for somatic embryo genesis induction. Remove the

most apical expanding leaves (8–12 mm) from proliferating

shoots and place them (abaxial side down) in test tubes ( see

Note 8 ) containing the IM medium. With a scalpel make 4–6

transverse cuts in the central part of the leaves ( see Note 11 ).

10. Transfer the cultures to a growth chamber, in the dark, at

25 °C. Somatic embryo s start to appear after 6–8 weeks of

culture (Fig. 3a ). About 2 weeks later, globular somatic

embryo s are formed (Fig. 3b ) ( see Note 12 ).

11. Transfer 2–4 mature cotyledonary somatic embryo s (Fig. 3c )

per test tube containing 15 mL of GM and keep the cultures

under a 16 h photoperiod of 15–20 μmol/m

2 /s (cool-white

ﬂ uorescent lamps) at 25 °C ( see Note 13 ). Germination can be

seen after 10–15 days (Fig. 3d ) of culture on GM medium.

12. Remove from the test tubes well-rooted ( see Note 14 ) plant-

lets (Fig. 3e ), measuring 3–4 cm in the aerial part, and wash

the roots with tepid tap water to get rid of agar ( see Note 15 ).

13. For acclimatization, transfer the plantlets to pots (250 mL vol-

ume) containing a mixture of sand and perlite (1:1), previously

sterilized by autoclaving.

14. Place the containers in a greenhouse under 18–20 °C until

plantlets reach 8–10 cm ( see Note 4 ).

João F. Martins et al.

335

Fig. 3 Somatic embryo formation and plant regeneration. ( a ) Early stages of somatic embryo development. ( b )

Somatic embryos mostly at the globular stage. ( c ) Cotyledonary embryo. ( d ) Germinated somatic embryos

after 12 days on GM medium. ( e ) Well-developed plantlets of somatic embryo origin just before to be potted.

( f ) Several plantlets obtained by somatic embryogenesis

Somatic Embryogenesis in Strawberry Tree

336

15. Transfer the plantlets (Fig. 3f ) to larger containers (1 L volume)

containing a mixture of autoclaved substrate, composed of

sand and peat (1:1).

16. When the plants reach 30–40 cm, transfer them to ﬁ eld condi-

tions. Figure 4 summarizes the different steps of the methodol-

ogy used to achieve plant regeneration through this protocol.

4 Notes

1. Branches must be healthy and from the upper part of the plant.

Avoid using older woody branches. These branches sprout as

well as younger branches, but the in vitro response of the

explants is lower. Spring is the best time to initiate the cultures,

but it was found that branches collected during other periods

of the year can also be used without signiﬁ cant behavioral

differences.

2. Recent data (unpublished) showed that the pH of the culture

media strongly inﬂ uences the in vitro response and that this

effect is genotype dependent. Thus, it may be necessary to use

different pH values for different trees but this has to be estab-

lished experimentally.

3. Spraying (three times a week) the branches with a 9.0 μM solu-

tion of BA stimulates epicormic shoot development ( see ref.

11 ) but is not absolutely necessary to induce it. Furthermore,

it can affect further response of the apices and nodal segments

in culture by reducing shot growth.

4. Covering the branches with polypropylene plastic bags main-

tains a higher humidity and may help to stimulate epicormic

shoot development.

5. Although both shoot apices and nodal segments can be used

for axillary shoot proliferation, the ﬁ rst gave better rates of

proliferation.

6. Strawberry tree tissues are very sensitive to ethanol. Do not keep

the explants in contact with this alcohol longer than suggested.

7. As a general procedure, we submit the explants to calcium

hypochlorite for 15 min. The effect of this treatment can be

genotype dependent, e.g., leaves of some trees are more sensi-

tive than others. If the tissues start to bleach before the indi-

cated time, immediately remove the explants and wash them.

8. Other containers such as Magenta boxes or glass boxes can be

used instead of test tubes. From the different containers tested,

it was found that test tubes covered with plastic caps are the

most effective for shoot proliferation. For somatic embryo gen-

esis induction, leaves can also be cultured in Petri dishes or

other plastic or glass containers.

João F. Martins et al.

337

Fig. 4 General overview of the process of plant regeneration of Arbutus unedo through somatic

embryo genesis

Somatic Embryogenesis in Strawberry Tree

338

9. More than one explant can be placed per test tube; however,

this way the probability of contamination is higher.

10. In general, using this protocol, A. unedo tissues do not release

phenolics to the culture medium. However, we have found

phenol exudation and oxidation when working with some gen-

otypes. In this case tissue cultures must be placed in a growth

chamber at 25 ± 1 °C under dark conditions for a week and

then transferred to light conditions. The inclusion of 100 mg/L

ascorbic acid in the medium also helps to reduce phenol oxida-

tion and necrosis of the explants.

11. Somatic embryo genesis can be achieved even without wound-

ing the leaves. However, this procedure increases the rates of

somatic embryo genesis induction.

12. Somatic embryo formation is asynchronous and embryos at

different developmental stages, from globular to cotyledonar

stage, can be seen in the same explant. The rates of somatic

embryo genesis induction are quite variable among different

trees. In the tested conditions, recalcitrant genotypes as well as

high responsive genotypes (i.e., with over 50 % of induction

rates) were found. The number of somatic embryos per

embryogenic explants is also variable, with some explants giv-

ing rise to only a few embryos, whereas others produce over

100.

13. Do not transfer to the germination medium the non-

cotyledonar embryos. They will not germinate or will undergo

precocious germination. Try to select morphologically normal

embryos since off-type embryos usually give rise to abnormal

plantlets.

14. Plantlets are considered well-rooted when showing at least two

well-developed roots. In some cases, only the stem part of the

plantlets develops after somatic embryo germination . In these

cases, the shoots can be rooted following an IBA (indole-3-

butyric acid) treatment followed by transfer to an auxin-free

medium, as described in ref.

9 .

15. Roots must be washed carefully not to break them, thus pre-

venting hindrance of the success of the next steps.

Acknowledgments

This work was supported by Fundação para a Ciência e Tecnologia

(Portugal), research contract PTDC/AGR-FOR/3746/2012,

and by a postdoctoral research fellowship (SFRH/

BPD/91461/2012) awarded to Sandra Correia.

João F. Martins et al.

339

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(2005) Propagation of the strawberry tree

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trees of Arbutus unedo L. (strawberry tree). N

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_15, © Springer Science+Business Media New York 2016

Chapter 15

Somatic Embryogenesis in Olive ( Olea europaea

L. subsp. europaea var. sativa and var. sylvestris )

Eddo Rugini and Cristian Silvestri

Abstract

Protocols for olive somatic embryogenesis from zygotic embryos and mature tissues have been described

for both Olea europaea sub. europaea var. sativa and var. sylvestris . Immature zygotic embryos (no more

than 75 days old), used after fruit collection or stored at 12–14 °C for 2–3 months, are the best responsive

explants and very slightly genotype dependent, and one single protocol can be effective for a wide range

of genotypes. On the contrary, protocols for mature zygotic embryos and for mature tissue of cultivars are

often genotype speciﬁ c, so that they may require many adjustments according to genotypes. The use of

thidiazuron and cefotaxime seems to be an important trigger for induction phase particularly for tissues

derived from cultivars. Up to now, however, the application of this technique for large-scale propagation

is hampered also by the low rate of embryo germination; it proves nonetheless very useful for genetic

improvement.

Key words Double regeneration technique , Immature and mature zygotic embryos , Mature tissues ,

Olea europaea L. var. sativa , Olea europaea L. var. sylvestris , Somatic embryogenesis

1 Introduction

Until recent years, olive breeding was limited, mainly due to the

long juvenile phase of the seedlings and to the scarce knowledge

related to genetic aspects, which discouraged breeders and public

researchers. For this reason, biotechnology-mediated genetic

improvement, which still requires efﬁ cient in vitro regeneration

protocols from cells or tissues, is a precious tool in support of con-

ventional breeding. Actually, it is possible to easily regenerate

somatic embryos or adventitious shoots from juvenile material,

such as zygotic embryos, due to the high morphogenetic ability of

tissues collected from immature or mature seeds [

1 – 10 ] in both

Olea europaea var. sativa and Olea europaea var. sylvestris. However,

these structures cannot be employed either for plant cloning or

for breeding as, due to the high heterozygosis of the Olea spp.,

342

they produce plants genetically different from the mother plant.

On the other hand, somatic embryogenesis is very difﬁ cult to

obtain from mature tissues of cultivars; moreover, this phenome-

non is strongly genotype dependent. In corroboration of that, up

to now somatic embryogenesis has been successfully reported only

in the cultivars Canino, Moraiolo [

11 ], and Dahbia [ 12 , 13 ] of O.

europaea L. var. sativa and in one genotype of O. europaea L. var.

sylvestris [

14 ]. The var. sylvestris has been taken into account in this

chapter because it may contribute to improve the cultivated variet-

ies of O. europaea L. var. sativa , since it belongs to the primary

gene pool [

15 ] of the genus. In our experience [ 11 ], regeneration

of adventitious buds, which normally provokes rejuvenation of the

regenerants, seems to be one of the key factors to obtain somatic

embryogenesis from mature tissues of cultivars, together with the

employment of a proper combination of basal medium and plant

growth regulators (PGRs). However, the results obtained in the cv.

Dahbia demonstrated that rejuvenation through adventitious

shoot organogenesis is not essential for the induction of embryo-

genic response [

13 ]. Regarding the PGRs, thidiazuron (TDZ)

seems to be an important trigger for the induction phase, since it

was used successfully for somatic embryogenesis induction.

Recently, the development of early stages of embryogenic cell sus-

pension culture from mature olive leaf-derived calli of the cv.

Chetoui has also been reported, but the development of well-

formed embryos was not achieved [

16 ].

The ability of somatic embryos to form secondary embryos

and, consequently, a cyclic embryogenesis can both be useful in the

unconventional breeding of olive, although the derived plants

acquire juvenility over time and, hence, delay their adult phase

[

17 ]. Concerning somaclonal variation, in our experience, an evi-

dent phenotypic variability was not observed yet, after many years

of in-ﬁ eld observation of plants of cv. Canino derived from somatic

embryogenesis (Fig.

1f–i ). This fact was conﬁ rmed by Lopes et al.

[

18 ] in Olea spp., where the genome integrities have been main-

tained throughout the embryogenesis process. On the contrary, a

report described a different vegetative behavior (bushy and colum-

nar phenotype) in plants derived from somatic embryos, originated

from one cotyledon of the cv. Frangivento [

19 ]. These conﬂ icting

results suggest to pay attention to the use of somatic embryogen-

esis for propagating true-to-type olive plants. On the other hand,

the high multiplication potential of cyclic somatic embryogenesis,

originated from mature tissues of elite cultivars, makes it a very

suitable technique to induce somaclonal variation under selective

pressure of biotic and abiotic stresses, as well as to be applied in

genetic transformation for the introduction of some agronomical

useful genes, as reviewed by Rugini et al. [

17 ].

Eddo Rugini and Cristian Silvestri

343

2 Materials

1. Material used to induce embryogenic callus: (1) Immature

olive fruits harvested between 30 and 75 days after blooming,

to be used both at harvest time or after storage at 12–15 °C

for 2–3 months, before extracting the embryos; (2) mature

olive fruits of O. europaea L. var. sativa and of O. europaea L.

var. sylvestris ; (3) in vitro shoots from mature trees of the cv.

Canino, Moraiolo, or Dahbia, micropropagated on OM

medium (Table

1 ; see Note 1 ) supplemented with 2190 mg/L

L - glutamine, 1 mg/L zeatin riboside (ZR), 3.6 % mannitol (or

3 % sucrose), and 0.6 % agar; (4) adventitious buds regener-

ated from leaf petioles of the cv. Canino or Moraiolo; and (5)

young plants of O. europaea L. var. sylvestris (about 2-year-old

plants, grown in greenhouse and derived from ﬁ eld plants of

over 30 years of age) used to collect young leaves.

Fig 1 Somatic embryogenesis by using a double regeneration system from in vitro growing shoot (cv. Canino).

( a ) Adventitious bud petioles from in vitro growing shoots. ( b ) Somatic embryogenesis from leaﬂ ets of a small

adventitious bud that was previously regenerated from the petiole. ( c ) Secondary somatic embryogenesis from

epidermal cell layer of somatic embryos and teratoma (cyclic somatic embryogenesis). ( d ) Histology of sec-

ondary somatic embryogenesis from epidermal cell layer of somatic embryos. ( e ) Embryo conversion. ( f )

14-year- old olive from somatic embryogenesis. ( g ) View of 7-year-old in-ﬁ eld plants derived from somatic

embryos. ( h ) Comparison of young branches with leaves and drupes collected from somatic embryo-derived

plants and the donor plant. ( i ) Detail of leaves and fruits from previous photos. Up to now, no evident variation

has been observed in vegetative and reproductive habits

Somatic Embryogenesis in Olive

344

2. Deionized water and sterile water.

3. Commercial bleaching solution.

4. 70 % ethanol.

5. MS medium [

20 ], OM medium [ 21 ], zeatin riboside (ZR),

benzylaminopurine (BAP), 3-indolbutyric acid (IBA),

α-naphthaleneacetic acid (NAA), 6-γ-γ-(dimethylallylamino)-

purine (2-iP), TDZ, Difco Bacto agar, plant agar, Tween 20.

6. Equipment: Laminar ﬂ ow hood, growth chamber, dissecting

microscope, pH meter, and autoclave.

Table 1

Rugini olive medium composition (as reported by Duchefa, the Netherlands)

Micro elements ( mg/L )

CoCl

2 · 6H 2 O 0.025

CuSO

4 · 5H 2 O 0.250

FeNaEDTA 36.700

H

3 BO

3 12.400

KI 0.830

MnSO

4 · H 2 O 16.900

Na

2 MoO

4 · 2H 2 O 0.250

ZnSO

4 · 7H 2 O 14.300

Macroelements ( mg/L )

CaCl

2 332.16

Ca(NO

3 )

2 416.92

KCl 500.00

KH

2 PO

4 340.00

KNO

3 1100.00

MgSO

4 732.60

NH

4 NO

3 412.00

Vitamins ( mg/L )

Biotin 0.05

Folic Acid 0.50

Glycine 2.00

Myoinositol 100.00

Nicotinic acid 5.00

Pyridoxine HCl 0.50

Thiamine HCl 0.50

Eddo Rugini and Cristian Silvestri

345

7. Disposables: Petri dishes (25 mm × 90 mm), 25-well multiwell

plates, pipettes, Whatman 3 mm ﬁ lter paper, 0.22 μm Millipore

ﬁ lters, forceps, scalpels, Paraﬁ lm, and Jiffy pots.

3 Methods

1. Adjust the pH of the media at 5.8 with 0.1 N KOH or 0.1 N

HCl and solidify with plant agar (unless otherwise stated).

2. Sterilize all the culture media by autoclaving at 121 °C for

20 min.

3. Add the ﬁ lter-sterilized hormones after autoclaving.

1. Break the stones and remove the seeds from the stony

endocarp.

2. Surface sterilize the seeds with 10 % commercial bleaching

solution for 10–15 min.

3. Rinse three times the seeds with sterile water.

4. Soak the seeds for at least 24 h in sterile water at room

temperature.

5. With a scalpel, remove the zygotic embryos by either a longi-

tudinal or transversal cut across the seed teguments and

endosperm.

6. Place the embryos [

10 – 15 ] horizontally in each Petri dish,

containing 20 mL of half-strength MS medium [

20 ] supple-

mented with 0.1–0.5 mg/L BAP, 2 % sucrose, and 0.6 % Difco

Bacto agar; seal the dishes with Paraﬁ lm.

7. Keep the cultures in the dark at 23 ± 1 °C and subculture them

in fresh medium every 30 days.

8. Subculture the embryogenic calli ( see Note 2 ) in the same

medium reducing BAP concentration at 0.05 mg/L or in

hormone- free medium.

1. Break the stones and remove the seeds from the stony

endocarp.

2. Sterilize the seeds for 1 min with 70 % ethanol, and then soak

them for 20 min in 10 % commercial bleaching solution plus

20 drops/L of Tween 20.

3. Rinse three times with sterile distilled water, and soak for 24 h

in sterile distilled water in the dark, at 24 ± 1 °C.

4. Extract the embryos and dissect them in three parts: proximal

and distal half of cotyledons and radicle ( see Note 3 ).

5. Transfer the explants to Petri dishes containing a modiﬁ ed

OM medium (OMc, see Note 4 ) with 0.5 mg/L 2-iP, 5 mg/L

IBA, and 0.6 % Difco Bacto agar.

3.1 Somatic

Embryogenesis

from Immature Zygotic

Embryos

3.2 Somatic

Embryogenesis

from Mature Zygotic

Embryo of Olea

europaea var. sativa

and var. sylvestris

Somatic Embryogenesis in Olive

346

6. Maintain the Petri dishes in 16 h photoperiod, under

40 μmol/m

2 /s of light intensity, at 24 ± 1 °C, for 21 days.

7. Transfer 25 callusing explants after 14–21 days to Petri dishes

containing gelled 20 mL hormone-free OMc medium, at the

abovementioned environmental conditions, to induce somatic

embryogenesis ( see Notes 5 and 6 ).

1. Collect leaf petioles from shoots, micropropagated on OM

medium supplemented with 1 mg/L ZR, 3.6 % mannitol (or

3 % sucrose), and 0.6 % agar.

2. Place the leaf petioles in Petri dishes containing 20 mL MS

medium supplemented with TDZ, at concentration ranging

between 2 and 10 mg/L, 2 % sucrose, 0.6 % Difco Bacto agar

( see Note 7 ), and seal with Paraﬁ lm with the aim of regenerat-

ing adventitious buds (Fig.

1a ).

3. Dissect the leaﬂ ets from the neo-formed adventitious buds

when they are no longer than 1–3 mm. Place them individu-

ally in a 25 multiwell plate (Sterilin) with 3 mL of OMc

medium supplemented with 1 mg/L hydrolyzed casein, 3 %

sucrose, 200 mg/L cefotaxime, 0.1 mg/L BAP, 0.1 mg/L

2-iP, 0.5 mg/L IBA, and 0.6 % Difco Bacto agar.

4. Store then the cultures in the dark at 24 ± 1 °C.

5. After about 1 month, when morphogenetic masses are pro-

duced from petioles, transfer them to a ﬁ lter paper (Whatman

no. 3) of the same diameter of the Petri dishes, with 5 mL

OMc liquid medium.

6. Every 3 weeks, remove the exhausted medium by pipetting and

replace it with an equivalent volume of fresh liquid medium,

until masses with pro-embryonic structures appear (they are

recognizable by a small spherical callus, with diameter of

1–2 mm and a smoothly yellowish surface; the masses continue

to enlarge in size and start to differentiate somatic embryos).

7. Place the neo-formed embryos (Fig.

1b ) on hormone-free

OMc medium with an agar content reduced to a value of 0.3 %

( see Note 8 ).

1. Collect the young leaﬂ ets and petioles from in vitro micro-

propagated shoots, growing on OM medium supplemented

with 1 mg/L ZR, 3.0 % sucrose, and 0.6 % agar.

2. Place the explants in Petri dishes on a shaker (regulated at

60 rpm), ﬁ lled with liquid induction medium, consisting of

half-strength MS medium supplemented with 6.60 mg/L

TDZ and 0.1 mg/L NAA, for 4 days.

3. Transfer the explants to hormone-free half-strength MS

medium, solidiﬁ ed with 0.6 % Difco Bacto agar for 8 weeks.

3.3 Somatic

Embryogenesis

from Mature Tissue

Explants of In Vitro

Grown Shoots

3.3.1 Cv. Canino

and Moraiolo

3.3.2 Cv. Dahbia

Eddo Rugini and Cristian Silvestri

347

4. Transfer the explants to the expression ECO medium

( see Note 9 ), supplemented with 0.1 mg/L BAP, 0.1 mg/L

2-iP, 0.05 IBA, and 0.6 % Difco Bacto agar.

1. Collect young leaves formed on 2-year-old potted plants

grown in greenhouse.

2. Sterilize the leaves for 1 min in 70 % ethanol, followed by

10 min immersion in 25 % commercial bleaching solution.

3. Rinse three times with sterile distilled water.

4. Dissect the leaves and place 10 explants (petioles, median, and

distal portion of leaf blades) in Petri dishes, containing MS

medium supplemented with 3 % sucrose, 1 mg/L ZR,

2.5 mg/L IBA, and 0.7 % Difco Bacto agar ( see Note 10 ).

5. Store the cultures in the dark at 22 ± 1 °C for 3 months.

6. Transfer the cultures on hormone-free MS medium in order

to achieve somatic embryogenesis expression.

7. Subculture the tissue explants every 4 weeks in the same

medium to maintain somatic embryogenesis expression.

1. Separate embryos from calli or from original tissues.

2. Place 5–6 embryos in 6-well multiwells (Sterilin) with 2 mL

OMc liquid medium, supplemented with 0.3 mg/L ZR; place

the multiwells on a gyratory shaker at 80 rpm.

3. Place the cultures in a growth chamber at 16 h light photope-

riod (40 μmol/m

2 /s).

4. Transplant the young plantlets with 2–3 pairs of leaves to Jiffy

pots for the acclimatization.

5. Transfer them to the greenhouse under high relative humidity

(80 %), at a temperature of about 23 °C and gradually reduce

the humidity.

4 Notes

1. Olive medium [ 21 ], commercialized in powder form by the

company DUCHEFA, is normally used for long-term prolif-

eration of olive shoots with 35-day subcultures. Olive medium

is characterized by a high content of Ca, Mg, S, Cu, and Zn

(Table

1 ). This medium usually induces tender shoots, with less

basal callus, when subcultures are done regularly every month.

2. The embryos will be visible after 5–6 weeks in culture.

Usually, the percentage of zygotic embryos forming somatic

embryos directly from the tissues (direct somatic embryogen-

esis) is considerably higher than embryos originated from the

neo-formed calli (indirect somatic embryogenesis), which

lose their embryogenic ability after one to two subcultures.

3.4 Somatic

Embryogenesis

from Mature Tissue

Explants of In Vivo

Grown Plants of Olea

europaea var. sylvestris

3.5 Embryo

Maturation

and Conversion

to Plantlets

Somatic Embryogenesis in Olive

348

Secondary embryogenesis normally takes place from epidermal

tissues of neo-formed embryos or teratoma, which can be

maintained for several subcultures in the same medium,

allowing cyclic somatic embryogenesis.

3. The radicles show a somatic embryogenesis ability higher than

that of cotyledon explants.

4. Modiﬁ ed OMc medium is a medium in which OM macroele-

ments are replaced with those of BN [

22 ], and the organic

compounds (myoinositol, glycine, thiamine HCl, pyridoxine

HCl, nicotinic acid, biotin, folic acid) are ten times higher

than the ones of OM medium.

5. Embryo formation, although in lower percentage than in

hormone- free medium, can be achieved also in modiﬁ ed OMc

medium, supplemented with 0.5 mg/L IBA [

23 ].

6. For Olea europaea L. var. sylvestris , transfer the explants to hor-

mone-free OMc medium or supplemented with 0.1–1 mg/L

IBA after 21 days, in order to get somatic embryos.

7. Rugini and Mencuccini [

24 ] obtained shoot organogenesis by

using both TDZ alone (at concentrations ranging between 2

and 10 mg/L) and in combination with auxin (2 mg/L TDZ

plus 0.5 mg/L NAA).

8. Secondary somatic embryos (Fig.

1c ) originate from the epi-

dermis, or rarely from the ﬁ rst subepidermal layer, of the

embryos (Fig.

1d ) [ 25 ]. A limited number of cells of the pri-

mary explant are apparently involved in the formation of

somatic embryos [

26 ]. Embryogenic masses and normal

embryos produce various embryo types, consisting of single

embryos of normal shape, single embryos with developed cot-

yledons, single embryos with more than two cotyledons, and

double or multiple embryos fused together [

9 ]. Activated

charcoal at 0.1 % can help the production of cyclic secondary

embryogenesis, originating from normal and abnormal

embryos or embryogenic masses.

9. ECO medium [

8 ] consists of ¼ OM macroelements, ¼ MS

microelements, and ½ OM vitamins supplemented with

550 mg/L

L -glutamine. Embryo “germination” (i.e., the

conversion of embryos to plantlets, Fig.

1e ) is difﬁ cult,

although the hypocotyl elongation is usually successful, while

epicotyl development does not always occur, due to the abnor-

mal embryo structure. However, 1 week at cold treatment

(4 °C) often increases embryo germination.

10. Somatic embryogenesis from petioles collected from in vivo

grown plants has also been achieved, although in low efﬁ -

ciency [

14 ], in the medium proposed by Rugini and Caricato

[

11 ], consisting of OM medium supplemented with 0.1 mg/L

BAP, 0.05 mg/L IBA, and 0.1 mg/L 2iP.

Eddo Rugini and Cristian Silvestri

349

References

1. Mazri MA, Belkoura I, Pliego-Alfaro F,

Belkoura M (2012) Embryogenic capacity of

embryo-derived explants from different olive

cultivars. Acta Hortic 929:397–403

2. Rugini E, Tarini P (1986) Somatic embryo-

genesis in olive ( Olea europaea L.). In:

MoetHennessy (ed) Proceedings of the

Conference in Fruit Tree Biotechnology,

Paris. p 62

3. Shibli RA, Shatnawi M, Abu E, Al-Juboory

KH (2001) Somatic embryogenesis and plant

recovery from callus of “Nabali” olive ( Olea

europaea L.). Sci Hortic 88:243–256

4. Maalej M, Drira N, Chaari-Rkhis A, Trigui A

(2002) Preliminary results of somatic embryo-

genesis from young zygotic embryos of olive

tree. Acta Hortic 586:899–902

5. Cerezo S, Mercado JA, Pliego-Alfaro F (2011)

An efﬁ cient regeneration system via somatic

embryogenesis in olive. Plant Cell Tiss Org

106(2):337–344

6. Maalej M, Chaari-Rkhiss A, Drira N (2006)

Contribution to the improvement of olive tree

somatic embryogenesis by mineral and organic

analysis of zygotic embryos. Euphytica

151(1):31–37

7. Rugini E (1988) Somatic embryogenesis and

plant regeneration in olive ( Olea europaea L.).

Plant Cell Tiss Org 14:207–214

8. Cañas LA, Benbadis A (1988) Plant regenera-

tion from cotyledon fragments of the olive tree

( Olea europaea L.). Plant Sci 54:65–74

9. Peyvandi M, Ebnrahimzadeh H, Majd A

(2008) Somatic embryos at different maturity

stages in two olive cultivars. Acta Hortic

791:213–216

10. Orinos T, Mitrakos K (1991) Rhizogenesis

and somatic embryogenesis in calli from wild

olive ( Olea europaea var. sylvestri s (Miller)

Lehr) mature zygotic embryos. Plant Cell Tiss

Org 27:183–187

11. Rugini E, Caricato G (1995) Somatic embryo-

genesis and plant recovery from mature tissues

of olive cultivars ( Olea europaea L.) “Canino”

and “Moraiolo”. Plant Cell Rep 14(4):

257–260

12.

Mazri MA, Elbakkali A, Belkoura M, Belkoura I

(2011) Embryogenic competence of calli and

embryos regeneration from various explants of

Dahbia cv, a moroccan olive tree ( Olea europaea

L.). Afr J Biotechnol 10(82):19089–19095

13. Mazri MA, Belkoura I, Pliego-Alfaro F,

Belkoura M (2013) Somatic embryogenesis

from leaf and petiole explants of the moroccan

olive cultivar Dahbia. Sci Hortic 159:88–95

14. Capelo AM, Silva S, Brito G, Santos C (2010)

Somatic embryogenesis induction in leaves

and petioles of a mature wild olive. Plant Cell

Tiss Org 103(2):237–242

15. Rugini E, De Pace C, Gutierrez-Pesce P,

Muleo R (2011) Olea. In: Kole C (ed) Wild

crop relatives: genomic and breeding resources

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Msellem M, Ghezel R, Bouzid S (2011)

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Olea spp. using nuclear microsatellites. Trees

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473–497

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olive ( Olea europaea sativa L.) cultivars with

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shoots and embryos. Sci Hortic 24:123–134

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Somatic Embryogenesis in Olive

351

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_16, © Springer Science+Business Media New York 2016

Chapter 16

Somatic Embryogenesis in Crocus sativus L.

Basar Sevindik and Yesim Yalcin Mendi

Abstract

Saffron ( Crocus sativus L.) is one of the most important species in Crocus genus because of its effective

usage. It is not only a very expensive spice, but it has also a big ornamental plant potential. Crocus species

are propagated by corm and seed, and male sterility is the most important problem of this species. Hence,

somatic embryogenesis can be regarded as a strategic tool for the multiplication of saffron plants. In this

chapter, the production of saffron corms via somatic embryogenesis is described.

Key words Crocus sativus L , Ornamental saffron , Somatic embryo genesis , Tissue culture

1 Introduction

The Iridaceae family contains 65 genus and 2025 species. It has a

big ornamental potential because of its leaf and ﬂ ower characteris-

tics [

1 ] . Generally, this family includes geophytes and monocotyle-

don [

2 ]. Crocus is the most important genus of the Iridaceae family.

It includes plants with rhizomes, corms, and bulbs . This genus

expands in tropical and subtropical regions of the Northern

Hemisphere, particularly in Southeastern Europe, North Africa,

and temperate Asia (Western and Central Asia) [

3 – 5 ] . Many coun-

tries, such as Iran, India, Greece, Spain, Italy, Turkey, France,

Switzerland, Israel, Pakistan, Azerbaijan, China, Egypt, the United

Arab Emirates, Japan, Afghanistan, and Iraq (and, recently, also

Australia in the Southern Hemisphere), produce saffron [

4 ].

Crocus has invariably been cultivated by means of traditional labor-

intensive methods which contribute to its very high price.

Crocus belongs to subfamily Crocoideae , one of the most

crowded subfamily of the Iridaceae family [

6 ]. The genus includes

nearly 100 species which are commonly used as popular orna-

mental plants [

7 ]. All the members of the genus are geophytes

and perennial plant species with an underground storage organ

and renewable buds. They are generally propagated not only by

352

seeds but also by specialized underground storage organs, such as

bulbs , corms, tubers, or rhizomes [

8 ]. Crocus is highly prized as

garden plants for their colorful ﬂ owers and as horticultural variet-

ies for industrial applications. Crocus genus includes native spe-

cies, especially in Greece, Turkey, Iran, and India, while other

countries (Italy, Hungary, and Spain) have representatives of

Crocus species [

6 ]. This genus presents a wide variety of chromo-

some numbers (2 n = 6, 8, 10, 12, 14, 16, 18, 20, 22, 23, 24, 26,

28, 30, 32, 34, 44, 48) [

9 ].

Crocus sativus L., commonly known as saffron , is the most

common cultivated plant, used for different purposes (ornamen-

tal, medicinal, and as spice). It is unknown as a wild plant, repre-

senting a sterile triploid derived from the naturally occurring

diploid C. cartwrightianus Herbert. Some archaeological and his-

torical studies indicate that domestication of saffron dates back to

2000–1500 years B.C. [

4 ]. Presently, saffron is commonly culti-

vated for its stigmas (used dried to produce the spice), while other

parts of the plant (such as leaves or petals) are useless; moreover,

a toxic effect of the bulbs on animals has been described [

10 ].

Because of its autotriploidy, saffron is multiplicated through the

formation of daughter corms from the mother corm, and its

breeding is very difﬁ cult. The saffron plant produces about 160

compounds, and crocin, safanal, picrocrocin, and crocetin are the

most valuable. Crocin, typically deep red in color, quickly dis-

solves in water to form an orange-colored solution, thereby mak-

ing crocin widely used as a natural food colorant. The second

most abundant component is picrocrocin that gives the taste of

saffron [

11 ]. These components provide different usage both in

medicine industry and in alternative medicine. Recently, several

studies showed the anticarcinogenic and antitumor activity of saf-

fron. As ornamentals, although Crocus species have ﬂ owers with

different shapes and colors and different plant morphology, as

well, they are still not exploited in gardening as it is in their

potential.

Tissue culture is an effective method for producing plants from

species with propagation problems. Tissue culture provides many

beneﬁ ts for rapid multiplication, and somatic embryo genesis is one

of the most useful technique, as it can produce high numbers of

plants from embryos originating from somatic cells of various tis-

sues and organs. Somatic embryo s can be also used for the produc-

tion of synthetic seeds, for genetic transformation and other

biotechnological applications. This chapter deals with the induc-

tion of somatic embryogenesis from C. sativus L. corms. It lists the

required equipment and describes a stepwise protocol useful to

sterilize explants and equipments, prepare somatic embryogenesis

medium, and induce somatic embryo maturation and conversion

to plantlets.

Basar Sevindik and Yesim Yalcin Mendi

353

2 Material

C. sativus L. is a perennial herb, and the plants used in this study

were collected in a small area in the Northwest of Turkey

(Safranbolu district, in the province of Karabük). Here, blooming

of saffron starts in October and goes on up to the middle of

November. Plants should be collected early in the morning in the

middle of October, during the ﬂ owering period. Plant material is

kept in pots until the use of explants for in vitro culture.

1. Vertical fume hood, laminar ﬂ ow cabinet.

2. Autoclave for sterilization.

3. pH meter.

4. Tissue culture facilities: magnetic stirrers, magnetic bars, for-

ceps, scalpels, and micropipettes.

1. Salts and vitamins from MS medium ([

12 ]; Table 1 ).

2. Sucrose .

3. Gelrite (Duchefa, NL).

4. 2,4-Dichlorophenoxyacetic acid (2,4-D).

5. N

6 -(2-isopentenyl)adenine (2iP).

(a) 1 N KOH solution.

(b) 1 N HCl solution.

6. 250 and 500 mL glass ﬂ asks.

7. 500 mL beakers.

8. 1000 and 500 mL cylinders.

9. Sterile Petri dishes (9 cm in diameter).

10. Magenta boxes (77 mm × 77 mm × 97 mm).

1. Tap water.

2. 0.1 % HgCl

2 solution.

3. 70 % ethyl alcohol (ETOH).

4. Distilled water.

5. 500 mL beakers.

6. 250 and 500 mL cylinders.

7. 20 % sodium hypochlorite (Domestos

® , commercial bleach

solution at 4.5 % NaOCl ).

1. Turf.

2. Perlite.

3. Vials (3.5 cm × 3.5 cm).

4. Fungicide (Captan 50 %).

2.1 Plant Material

2.2 Laboratory

Equipments

2.3 Preparation

of Culture Media

2.4 Explant

Sterilization

2.5 Acclimatization

of Plantlets

Embryogenesis in Saffron

354

3 Methods

1. To prepare the SEI medium, prepare the stock solutions of

macro- and microelements from MS medium . Alternatively,

use ready-to-use MS powder preparation (4.4 g/L) by Sigma-

Aldrich (M5519).

2. Prepare 2,4-D and 2iP stock solutions at 100 mg/100 mL

concentration by dissolving 2,4-D in 2–3 mL ETOH and 2iP

in 1 N NaOH.

3. Add 2 mL of 2,4-D stock solution to 1 L of MS medium to

have a ﬁ nal concentration of 2 mg/L, and 1 mL 2iP stock

solution to have a ﬁ nal concentration of 1 mg/L.

4. Add 30 g/L sucrose , add distilled water up to 1 L, optimize

the pH between 5.6 and 5.8 with 1 N KOH and 1 N HCl, and

add 4 g/L Gelrite (Duchefa, NL) ( see Note 1 ).

3.1 Preparation

of Somatic

Embryogenesis

Induction (SEI)

and Somatic Embryo

Maturation (SEM)

Media

Table 1

MS formulation [ 12 ]

Macronutrients mg/L

NH

4 NO

3 1650

KNO

3 1900

CaCl

2 × 2H 2 O 440

MgSO

4 × 7H 2 O 370

KH

2 PO

4 170

Micronutrients mg/L

H

3 BO

3 6.2

MnSO

4 × 1H 2 O 16.9

ZnSO

4 × 7H 2 O 10.6

KI 0.830

Na

2 MoO

4 × 2H 2 O 0.25

CuSO

4 × 5H 2 O 0.025

CoCl

2 × 6H 2 O 0.025

Amino acid and Vitamins mg/L

Glycine 2

Nicotinic acid 0.1

Thiamine HCl 0.5

Pyridoxine HCl 0.5

Basar Sevindik and Yesim Yalcin Mendi

355

5. For media sterilization, autoclave MS medium at 121 °C for

15 min and 1.05 atm pressure.

6. Pour the medium into the Petri dishes (diameter, 90 mm) in

the laminar ﬂ ow cabinet.

7. To prepare the SEM hormone-free medium, follow the same

procedure reported above, with the exception of steps 2 and 3 .

1. Before surface sterilization, tunics are separated from the

corms. Wash tunic-removed corms under tap water for 30 min

to eliminate soil traces from corm surface (Fig.

1a ).

2. Treat the corms with 0.1 % HgCl

2 for 30 min under the verti-

cal fume hood ( see Note 2 ), and then wash the corms with

sterile distilled water for 6–7 times.

3. Treat the explants with 70 % ETOH ( see Note 3 ) under the

laminar ﬂ ow cabinet for 1 min. Wash the explants for three or

four times with sterile distilled water.

4. Treat the explants with 20 % NaOCl solution in the laminar

ﬂ ow cabinet for 20 min ( see Note 4 ), and then wash them with

sterile distilled water for six or seven times (Fig.

1b ).

1. Under the laminar ﬂ ow cabinet, cut the explants into small

pieces of 4 mm, on average (Fig.

1c ).

2. Plate them on the SEI medium (four explants per Petri dish).

3. Culture the explants in the climate chamber, at 25 °C in dark-

ness (Fig.

1d–f ).

4. After 6 weeks, somatic embryo s are observed from both direct

and indirect somatic embryogenesis.

5. For maturation, transfer the somatic embryo s on the SIM

medium to get germination of embryos (Fig.

1g, h ).

1. Autoclave the 1:1 turf/perlite mix at 121 °C, 1.05 atm of

pressure, for 15 min; put the mix into vials (5.3 cm in

diameter).

2. Separate the microcorms from the hormone-free MS medium

and wash them under tap water.

3. Solubilize 1 g fungicide (Captan 50 %) in 1 L of distilled water.

4. Dip microcorms with emblings into the fungicide and put

them into the vials.

5. Put the vials into the climate chamber at 25 °C, photoperiod

16 h and about 38 μmol m

−2 s

−1 of light intensity; close the

vials with transparent nylon tarpaulin. Two weeks later, trans-

fer the plantlets in 500 cc pots, inside a greenhouse.

3.2 Explant

Sterilization

3.3 Induction

of Somatic

Embryogenesis

and Somatic Embryo

Maturation

3.4 Plantlet

Acclimatization

Embryogenesis in Saffron

356

4 Notes

1. Optimize the pH using 1 N KOH and 1 N HCl.

2. Be careful when managing HgCl

2 , a carcinogenic compound.

Work in the vertical fume hood, use the safety mask, and do not

Fig. 1 Different stages of somatic embryo genesis in saffron . ( a ) Removal of tunics from the corms. ( b ) Surface

sterilization of the corms. ( c ) Culture of the explants containing apical buds on MS medium . ( d ) Callus stage

(CP) in indirect somatic embryogenesis. ( e ) Differentiation of callus into somatic embryos from apical buds

(AB). ( f ) Different stages of somatic embryos (HF, heart stage; GF, globular stage). ( g and h ) Embling elongation

from somatic embryos

Basar Sevindik and Yesim Yalcin Mendi

357

breathe the solution. Weigh 1 g HgCl

2 , put in 1 L of distilled

water, and mix until dissolved.

3. Put 700 mL of ETOH into the cylinder measure and supple-

ment with 300 mL of distilled water.

4. Put 200 mL of commercial bleach solution (Domestos

® , con-

taining 4.5 % NaOCl ) into a cylinder measure and supplement

with 2–3 drops of Tween20 (nonionic detergent, Sigma-

Aldrich, USA) and add distilled water up to 1 L.

References

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Micropropagation of Iridaceae. Plant Cell Tiss

Organ Cult 97:1–19

2. Adamson RS (1925) On the anatomy of some

shrubby Iridaceae. Trans Roy Soc S Afr 13(2):

175–195

3. Wani BA, Mohiddin FA (2009) Micro-

propagation of Genus Crocus—a review. Afr

J Agric Res 4(13):1545–1548

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Embryogenesis in Saffron

359

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

vol. 1359, DOI 10.1007/978-1-4939-3061-6_17, © Springer Science+Business Media New York 2016

Chapter 17

Somatic Embryogenesis in Lisianthus ( Eustoma

russellianum Griseb.)

Barbara Ruffoni and Laura Bassolino

Abstract

Somatic embryogenesis is, for the main ﬂ oricultural crops, a promising system for commercial scale-up,

providing cloned material to be traded as seedlings.

Somatic embryos, having the contemporary presence of root apical meristem and shoot apical meri-

stem, can be readily acclimatized. For Lisianthus it is possible to induce embryogenic callus from leaf frag-

ments of selected genotypes and to obtain embryos either in agarized substrate or in liquid suspension

culture. The production of somatic embryos in liquid medium is high and can be modulated in order to

synchronize the cycle and the size of the neoformed structures. The possibility to use the liquid substrate

with high propagation rates reduces labor costs and could support the costs of eventual automation. In this

paper we report a stepwise protocol for somatic embryogenesis in the species Eustoma russellianum.

Key words Cell suspension , Conversion , 2,4-D , Lisianthus , Somatic embryo s , Somatic embryo

maturation

1 Introduction

Lisianthus ( Eustoma russellianum Griseb. or Eustoma grandiﬂ o-

rum (Raf.) Shinn.) belongs to Gentianaceae family and is a moder-

ately cold-tolerant annual or biennial plant native to the southern

part of the United States and Mexico [

1 ]. It is commonly known

as “Texas bluebell” and “prairie gentian.” Lisianthus gained popu-

larity in the international ﬂ ower market due to its roselike ﬂ owers,

excellent postharvest life (the cut inﬂ orescences typically have a

vase life of 3–6 weeks) [

2 ], and its attractive range of colors. In

nature, the phenology of the species provides for the initial devel-

opment of a rosette that grows very slowly during winter, stems

elongation in spring, and blooms in summer [

3 ]. The domesti-

cated varieties can be adapted for the production of ﬂ owers

throughout the year in protected cultivation. In recent decades,

breeders have developed a variety of cultivars with respect to many

traits such as uniform ﬂ owering (it can produce up to 20–40 ﬂ ow-

360

ers per plant), lack of rosetting, heat tolerance, ﬂ ower color, and

size and form, including double ﬂ owers [

4 ]. Currently, it is among

the top ten cut ﬂ owers in the international Holland market.

Eustoma is commonly propagated by seed or cutting. A large

number of seedlings can be produced by seed propagation, but the

quality is not uniform due to variations in ﬂ owering time, plant

height, and the number of ﬂ owers. In some cultivars with marginal

variegation, or doubled petals, the seedlings show a wide range of

variation because of their heterozygous character [

5 ]. Methods for

micropropagation of Eustoma have been developed by several

authors and shoots can be easily regenerated from stem and leaf.

The shoot regeneration from petals was studied by Ruffoni et al.

[

6 ] in relationship to the stage of the ﬂ ower maturation. Large-

scale multiplication of selected plants with superior characters is

possible for all the cultivars showing for this trait poor genotype

variation [

7 ]. The effective multiplication rates depend on several

factors like genotype, culture media, plant growth regulator s

( PGR ), and type of explants [

8 – 10 ]. Somatic embryo genesis was

ﬁ rst reported in agarized substrate [

11 ] and, afterwards, in liquid

substrate [

12 ]. Attempts to obtain artiﬁ cial seed s after encapsula-

tion in alginate were also performed [

13 ], as well as the use of the

somatic embryo genesis protocol for genetic transformation [

14 ,

15 ]. Data about the productivity of several Lisianthus genotypes

were presented in 2006 [

16 ]; in the same year, somatic embryo-

genesis protocols were also reported by various authors [

16 , 17 ].

2 Materials

1. 70 % ethanol prepared with 70 mL 99.8 % ethanol and 29 mL

distilled water.

2. Aqueous NaOCl solutions of active chlorine (e.g., commercial

bleach ACE, Procter & Gamble, USA) plus two drops of sur-

face active agent Tween 20

® .

3. Autoclaved reverse osmosis water, 200 mL aliquots in 500-mL

culture vessels.

4. Magnetic stirrer, 1000-mL ﬂ ask or beaker.

5. Tissue culture facilities: Instruments (scalpel, forceps, spirit

burner to ﬂ ame sterilize instruments), laminar ﬂ ow hood, cul-

ture room.

6. Lisianthus potted plants as explant source.

1. Media based on the formulation of Murashige and Skoog (MS;

[

18 ]) for shoot propagation from apical and axillary buds of

young branches and root induction (Table

1 ).

2. Glass culture vessels (500 mL) with transparent caps.

2.1 Surface

Sterilization of In Vivo

Grown Source Material

2.2 Culture Media for

Micro propagation and

Callus Development

Barbara Ruffoni and Laura Bassolino

361

3. Petri dishes (9-cm diameter, 2-cm height Bibby Sterilin, Stone,

UK).

4. Paraﬁ lm (Paraﬁ lm

® M Barrier Film, SPI Supplies, West Chester,

USA).

5. Duran glass ﬂ asks or beakers, 1000-mL capacity (Schott AG,

Mainz, Germany).

1. 100, 250, 500 mL Erlenmeyer vessels with screw caps steril-

ized in autoclave at 121 °C, 1 atm (1.01325 bar) for 20 min.

2. Steel 500-μm mesh sieve (Fig.

1b ), combined with a beaker

having the same diameter (Fig.

1d ), enveloped together with

autoclavable plastic bag or paper, sterilized in autoclave as in

step 1 .

3. Sterile disposable pipette (5, 10, 25, 50 mL).

4. Fresh weight determination: Sterile ﬁ lter paper dishes, vacuum

pump, sterile ﬁ ltration device (all glass ﬁ lter holder, Millipore

® ).

5. Fluorescein diacetate test ( FDA [

19 ]): Non-sterile tubes,

Pasteur pipettes, inverse microscope with ﬂ uorescence equip-

ment (ﬁ lter set 09 ZEISS Co., l ex = 450–490 nm, l em = 520

nm).

6. Settled cell volume ( SCV ) determination: 100- and 250-mL

cylinders sterilized by autoclaving ( see step 1 ).

1. Plastic alveolary pots.

2. Potting medium consisting of peat (Trysubstrate Klasmann-

Deilmann) and sterilized sand (1:1, v/v).

2.3 Cell Suspension

Culture

2.4 Acclimatization

of Emblings to Ex Vitro

Conditions

Table 1

Chemical composition of the substrates for somatic embryo genesis in Lisianthus in agarized or

liquid medium (f.s., full strength)

A) Callus establishment

B) Cell

suspension

culture

C) Somatic embryos

development

D) Somatic

embryos

conversion

MS salts f.s. f.s. f.s. f.s.

MS vitamins f.s. f.s. f.s. f.s.

2,4-D 9.05 μM 9.05 μM – –

Kin – – – 1.5 μM

Sucrose 30 g/L 30 g/L 30 g/L 30 g/L

Technical agar 8 g/L – 8 g/L 8 g/L

pH 5.7 5.7 5.7 5.7

P.P.F.D. 35 μmol/m

2 /s 25 μmol/m

2 /s 35 μmol/m

2 /s

Lisianthus Embryogenesis

362

Fig. 1 ( a ) Lisianthus ﬂ ower branches; ( b ) leaf fragments washed and ﬁ ltered; ( c ) leaf debris and chlorophyll in

the washing medium; ( d ) ﬁ ltering apparatus, embryogenic cells synchronized for suspension culture; ( e )

somatic embryo s developed in agarized medium lacking in 2,4-D; ( f ) somatic embryo developed in liquid

hormone-free medium; ( g ) mature somatic embryo; ( h ) particular of the SAM of the somatic embryo (120×);

( i ) converted somatic embryos; ( l ) embling with true leaves ready for transfer to pot

Barbara Ruffoni and Laura Bassolino

363

3 Methods

1. Prepare 10 L of a 10× stock solution by dissolving 43 g of MS

(Duchefa Biochemie B.V., The Netherlands; cod M 0221.0010)

powder, containing a micro- and macro-element complex, in

9 L of deionized water. While stirring the water, add the pow-

der and stir until complete dissolution; bring the solution to a

ﬁ nal volume (10 L) by adding water; use 1 L of this solution

(4302.09 mg) for each liter of culture medium.

2. Prepare 250 mL of 1000x stock solution by dissolving MS

vitamins (Duchefa Biochemie B.V., The Netherlands; cod M

0409.0250) powder, containing 25.8 g mixed vitamins, in

deionized water, and stir until completely dissolved, eventually

warming the solution up to 30 °C. Use 1 mL vitamin stock

solution (103.1 mg) for each liter of culture medium.

3. Add any desired suitable supplement (commercial sucrose ,

usually 30 g/L) weighing the powder and dissolve by stirring.

4. PGR stocks: To prepare the two stock solutions at 1 mg/mL

of 2,4-dichlorophenoxyacetic acid (2,4-D) and kinetin (Kin),

or gibberellic acid (GA3), weigh 100 mg and add in a volu-

metric ﬂ ask 20–30 mL ethanol (99.8 %). Gently stir until com-

pletely dissolved and bring slowly the volume to 100 mL with

distilled water at room temperature. Store the stock solutions

as recommended (4 °C in light for 2,4-D and Kin). Add the

PGR as requested for each culture phase.

5. After adding all the components, while stirring, adjust the pH

of the medium to 5.7 by using NaOH 1 N or HCl 1 M.

6. Add 8 g/L technical agar (e.g., Duchefa Biochemie B.V., The

Netherlands); heat until clarity of the solution, stirring the

medium on an electric plate or heating in a microwave.

7. Dispense 62.5 mL medium each into 16 culture vessels (500 cc).

8. Sterilize the medium in autoclave at 121 °C, 1 atm (1.01325

bar) for 20 min. Allow medium to cool and solidify prior to

plant inoculation.

9. To pour culture medium in Petri dishes (9-cm diameter):

Follow instructions from point 1 to 5, add 8 g/L technical

agar , sterilize the medium in autoclave at 121 °C, 1 atm

(1.01325 bar) for 20 min, move the ﬂ ask (when cooled below

100 °C) under the laminar ﬂ ow hood, and dispense 25 mL

medium in each Petri dish to prepare 40 Petri dishes.

10. For liquid medium: Prepare the solution as previous step ( 1 –

5 ) without adding agar ; sterilize the medium in autoclave at

121 °C, 1 atm (1.01325 bar) for 20 min; move the ﬂ ask (when

cooled below 100 °C) under the laminar ﬂ ow hood. The liquid

media can be stored at 5 °C in the dark up to 30 days. Warm

the medium at room temperature (20 °C) before use.

3.1 Preparation

and Sterilization

of Culture Media

Lisianthus Embryogenesis

364

11. For FDA stock solution: Dissolve 5 mg of ﬂ uorescein diacetate

in 1 mL acetone, and store it at −18 °C. The stock solution can

be used for several months.

1. Excise young leaves of Lisianthus - selected genotypes from

mature greenhouse-grown plants (Fig.

1a ), maintained at 20

°C in 14-cm diameter pots, under natural light conditions.

2. Rinse explants in a detergent warm aqueous solution with a

few drops of liquid dish soap. Then, sterilize with 70 % ethanol

for 30 s, treat with NaOCl solution (1.75 % active chlorine) for

15 min, and rinse twice with sterilized distilled water.

3. Chop accurately the leaf tissue, wash the material and collect

the fragments over the steel ﬁ lter (Fig.

1b ), discard the liquid

with chlorophyll and debris (Fig.

1c ), and transfer the material

onto the medium for in vitro callus induction (Table

1, A ) for

30 days.

1. Transfer the callus formed after 30 days, observe with a bin-

ocular microscope, accurately cut off the brown leaf tissue, and

select the undifferentiated tissue. Then subculture the green

callus every 30 days.

2. Grow callus in growth chamber at the following conditions:

23 ± 1 °C and 16 h photoperiod (light intensity 35 μmol/m

2 /s

photosynthetic photon ﬂ ux density (PPFD)).

3. Transfer 1 g of the green friable callus, at 28–30-day intervals,

in fresh medium in Petri dish ( see Note 1 ).

1. Transfer 1 g of friable callus in 100-mL sterile Erlenmeyer ﬂ ask

and insert with a sterile pipette 25 mL of liquid embryogenic

medium (Table

1, B ).

2. Put the ﬂ ask in a gyratory shaker at 110 rpm in 16 h light pho-

toperiod at 35 μmol/m

2 /s PPFD ( see Note 2 ).

3. Every 15 days renew the medium: Transfer the ﬂ ask in the

sterile bench, let settle the cells for 15 min, gently aspire with

a sterile pipette the old liquid medium over the plant material

and discard, and then add the same amount of new liquid

medium previously stored for 1 h at room temperature.

4. Monitoring the growth by settled cell volume ( SCV ) ( see Note

3 ): Pour the cell suspension in a graduated cylinder of ade-

quate volume and allow the suspension to settle for 30 min

and record the volume of fraction occupied by the cells. Take

a second reading 30 min later; if the variation is higher than 5

%, take a third reading 30 min later.

5. Assessment of the cell viability, ﬂ uorescein diacetate test [

19 ]:

Dilute 1 mL of FDA stock solution with 2 mL of distilled

water in a test tube (it turns white milky). Then, mix 1 mL of

3.2 Callus

Establishment

3.3 Embryogenic

Callus Biomass

Proliferation

3.4 Embryogenic

Cell Suspension

Culture

Barbara Ruffoni and Laura Bassolino

365

the diluted FDA solution with 1 mL cell suspension and

incubated for 5 min at room temperature. Put onto a slide a

small amount of the solution containing FDA stained cells

(one or two drops) and observe it under a ﬂ uorescence

microscope by visualizing the greenish ﬂ uorescence of the

cells at 100–400× magniﬁ cation to calculate the percentage

of viable cells ( see Note 4 ).

6. Synchronization : After four subcultures pour out the suspen-

sion culture in the sieve (Fig.

1d ), the fraction that remains

over the sieve can be used for a second mother suspension cul-

ture; the fraction that pass through the grid contains cell

aggregates <500 μm; gently aspire this material and re-suspend

2 mL of it in a 100-mL Erlenmeyer ﬂ ask ﬁ lled with 30 mL of

fresh medium and put in agitation (Subheading

3.4 , step 2 )

( see Note 5 ).

7. Evaluation of the growth curve: Prepare 30 Erlenmeyer ﬂ asks

(100 mL) containing the same proportion of embryogenic

synchronized cell culture (2 mL) and fresh liquid medium (30

mL); put in agitation as described and evaluate fresh weight,

dry weight, and the pH of three samples separately every 2 days

( see Note 6 ).

1. In agarized medium: Transfer 500 mg of embryogenic callus

from medium A (Table

1 ) in Petri with medium C (Table 1 ) in

growth chamber at normal conditions ( see Subheading

3.3 ,

step 2 ); verify the somatic embryo development with binocu-

lar 25–30 days after transfer (Fig.

1e ).

2. In liquid phase: Transfer 1 mL of embryogenic synchronized

cell culture in medium C (Table

1 ) in Petri dishes ( see

Subheading

3.1 , step 9 ); store in light, in stationary phase at

23 ± 1 °C for 4 weeks ( see Note 7 ).

1. In agarized medium: 28–30 days after transfer in medium C

(Table

1 ), accurately select the neoformed structures from the

remaining callus and transfer them separately onto fresh

agarized medium D (Table

1 ) ( see Notes 8 and 9 ). Let them

grow for additional 30 days and then transfer to the green-

house (Subheading

3.7 , step 1 ).

2. In liquid medium: Filter the neoformed structures with a 500-

μm mesh sieve, gently collect the structures that remain in the

ﬁ lter (Fig.

1f–h ), and transfer them onto fresh agarized medium

D (Table

1 ). Let them grow for additional 30 days (Fig. 1i )

and then transfer to the greenhouse (Subheading

3.7 , step 1 ).

1. Transfer the plantlets with root and several true leaves (Fig.

1l ),

3.5-cm high, in alveolary pots prepared (Subheading

2.4 );

place for 21 days in the glasshouse with 70 % relative humidity,

3.5 Somatic Embryo

Development

3.6 Somatic Embryo

Conversion

and Growth

3.7 Embling

Acclimatization

Lisianthus Embryogenesis

366

maintained by mist system (10 s every 30 min); and then

decrease the humidity up to 60 %

2. The mean temperature can vary between 20 and 25 °C, and

for lighting, use natural light with 50 % shade provided by

polyester- aluminum net.

3. After 2 months transfer the acclimatized plants to 11-cm diam-

eter pots ﬁ lled with the same substrate.

4 Notes

1. Experiments were carried out in order to determine the best

2,4-D concentration to use for embryogenic callus produc-

tion. The primary callus was grown in the presence of 4.5,

9.05, or 18.10 μM 2,4-D for 30 days, in light ( see

Subheading

3.4 , step 2 ) or in dark at the same temperature.

The fresh weight of ten replications per each condition was

evaluated in grams and compared (Fig.

2 ). The ﬁ rst two con-

centrations gave similar results while the higher 2,4-D concen-

tration induced less amount of callus. Light conditions allowed

a better development of green and friable callus.

2. The presence of light during the cell culture increases the

embryogenic cell parameters as shown in Table

2 : Both viabil-

ity and SCV are low in darkness. The presence of a higher

amount of small clusters is also remarkable.

bb

a

bb

a

0

0.5

1

1.5

2

2.5

4.5 9.05 18.1

Callus growth (g)

2,4-D (μM)

light intensity 35 μmol m-2 s-1 P.P.F.D

Fig. 2 Evaluation of the growth of embryogenic callus (fresh weight), depending

on the presence or absence of light (16 h photoperiod at 35 μmol/m

2 /s PPFD). In

each light condition, different letters indicate means differing at p ≤ 0.05 by the

Student-Newman-Keuls test

Barbara Ruffoni and Laura Bassolino

367

3. The SCV value differs from packed cell volume ( PCV ) for the

sedimentation system; SCV uses gravity to sediment cells; PCV

is determined using a centrifuge, but in the case of embryo-

genic cells, the centrifugation increases the risk of bacterial

contamination without giving further evaluation elements on

the cell growth.

4. The milky solution of FDA is active for 15–20 min; after this

time the molecules crystallize and loose activity.

5. The mesh size is important to increase the cells and the cell

aggregates of a similar diameter; it was established that 500 μm

is a suitable grade that does not reduce the further somatic

embryo development (Table

3 ); in the same table, it is possible

to note that, at grade 200 μm, the number of somatic embryo

per mL signiﬁ cantly decreases in light conditions.

6. Growth curve of embryogenic cell culture: Starting from the

ﬁ ltered material ( see Subheading

3.4 , step 6 ), 30 Erlenmeyer

ﬂ asks were prepared with 3 mL of cells and 25 mL of medium B

Table 2

General features of the embryogenic cell cultures grown in dark or in the light (16 h photoperiod,

light intensity 35 μmol/m

2 /s PPFD). SCV , FDA test

Dark Light

Average viability ( FDA test) 85 % 92.5 %

Shape of single cells 80 % isodiametric <30 μm

20 % elongated Isodiametric <60 μm

Cell clusters (amount/type) ++/small +++/small and medium

Average SCV 15.38 % 18.3 %

Table 3

Mean number of somatic embryo s per milliliter of embryogenic cell

culture after transfer in 10 mL of liquid medium C (Table 1 ), in stationary

phase, in the dark, or in the light per 30 days, related to the cell size.

Filtration made by a steel sieve (clone LT3; 12 replications ± standard

error; mean separation by the Student-Newman-Keuls test; *, signiﬁ cant

at p ≤ 0.05, n.s., not signiﬁ cant)

Sieve mesh size

>500 μm <500 μm <200 μm p ( ≤0.05)

Light 16.7 ± 7.3b 13 ± 3.9ab 3.4 ± 1a *

Dark 8.5 ± 1.4 4.4 ± 1.6 11.7 ± 3.9 n.s.

Lisianthus Embryogenesis

368

(Table 1 ); at day 2, 4, 6, 8, 11, 13, and 16, three samples were

ﬁ ltered, the pH was detected, and the plant material was ﬁ rst

weighted (fresh weight) and then dried in oven at 80 °C for

36–48 h (dry weight). The growth curve (Fig.

3 ) shows a loga-

rithmic increase of both fresh and dry weight from day 6 to day

13, after which the growth decreases. The pH curve shows a

little variation and increasing values from day 8. A continuous

growth could be obtained by adding the fresh medium at day

10 during the log phase.

7. In Table

4 data are shown about the productivity of embryo-

genic cell cultures related to the 2,4-D concentration, demon-

strating that the concentration of 9 μM is the best to induce

somatic embryo s, and it also conﬁ rmed that in darkness the

embryogenic process is inhibited.

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

0.4

0.45

0

1

2

3

4

5

6

7

8

9

02468101316

fresh weight and pH

days

fresh weight (g)

pH

dry weight (g)

dry

weight

Fig. 3 Embryogenic suspension culture: growth curves of pH, fresh and dry

weight during culture in batch. Starting point (T0): synchronized cells ﬁ ltered at

500 μm

Table 4

Mean number of somatic embryo developed from suspension cultures

grown in the light (16 h photoperiod, light intensity 35 μmol/m

2 /s PPFD) or

in the dark and at different levels of 2,4-D after transfer in medium C

(Table 1 ) (clone LT3; 12 replications ± standard error; mean separation by

the Student-Newman–Keuls test;*, signiﬁ cant at p ≤ 0.05; n.s., not

signiﬁ cant)

2,4-D concentration (μM)

4.5 9.0 18.0 p

Light 0.83 ± 0.30c 27.00 ± 7.00a 7.00 ± 1.60b *

Dark 8.27 ± 2.25 10.00 ± 3.20 6.00 ± 1.70 n.s.

Barbara Ruffoni and Laura Bassolino

369

8. The somatic embryo productivity of cell suspension cultures of

several genotypes of Lisianthus from different origin has been

described by Ruffoni and Savona [16]. Interestingly, geno-

types suitable for pot plant production (from 3 to 22 somatic

embryos/mL of suspension culture) showed the lowest pro-

duction, while the clones suitable for cut ﬂ ower production

(up to 361 somatic embryos/mL of suspension) induced the

highest number of somatic embryos per milliliter, thus result-

ing in a possible development of 180,000 somatic embryos per

liter of culture.

9. “ Conversion ” is the term used for the germination of somatic

embryo s, the development of the cotyledonous leaves, and the

elongation of the root apex, occurring better in agarized

medium when Kin is added in a small amount (Table

5 ); the

liquid medium without hormones increases somatic embryo

hyperhydration , affecting the further development of the

structures in the greenhouse.

References

Table 5

Conversion percentage of the somatic embryo s developed in light

conditions after 35 days of culture in several germination substrates

(+ = presence; − = absence)

Conversion (%)

Liquid medium Agarized medium

H

2 O MS base MS base MS + GA

3 MS + Kin

SE converted into plant 32.0 39.0 5.6 29.5 51.5

Hyperhydration + − − − −

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Lisianthus russellianus hook. Acta Hortic

280:83–87

12. Massabò F, Ruffoni B (1996) Plant production

by somatic embryogenesis in cell suspension

cultures of Lisianthus russellianus hook. Plant

Tissue Cult Biotechnol 2(4):194–198

13. Ruffoni B, Massabò F, Giovannini A (1993)

Artiﬁ cial seeds technology in the ornamental

species Lisianthus and Genista . Acta Hortic

362:297–304

14. Semeria L, Vaira AM, Accotto GP, Allavena A

(1995) Genetic transformation of Eustoma

grandiﬂ orum Griseb. by microprojectile bom-

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Vaira AM, Accotto GP, Allavena A (1996)

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Tiss Org 47:67–72

16. Ruffoni B, Savona M (2006) Somatic embryo-

genesis in ﬂ oricultural crops: experiences of

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Cyclamen . In: Teixeira da Silva JA (ed)

Floriculture, ornamental and plant biotechnol-

ogy, vol 2. Global Science Books, Isleworth,

pp 305–313

17. Duong TN, Nguyen ST, Ngoc HM, Uyen PN,

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6(3):121–127

18. Murashige T, Skoog F (1962) A revised

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Barbara Ruffoni and Laura Bassolino

371

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 18

Somatic Embryogenesis in Two Orchid Genera

( Cymbidium , Dendrobium )

Jaime A. Teixeira da Silva and Budi Winarto

Abstract

The protocorm-like body (PLB) is the de facto somatic embryo in orchids. Here we describe detailed

protocols for two orchid genera (hybrid Cymbidium Twilight Moon ‘Day Light’ and Dendrobium

‘Jayakarta’, D . ‘Gradita 31’, and D. ‘Zahra FR 62’) for generating PLBs. These protocols will most likely

have to be tweaked for different cultivars as the response of orchids in vitro tends to be dependent on

genotype. In addition to primary somatic embryogenesis, secondary (or repetitive) somatic embryogenesis

is also described for both genera. The use of thin cell layers as a sensitive tissue assay is outlined for hybrid

Cymbidium while the protocol outlined is suitable for bioreactor culture of D. ‘Zahra FR 62’.

Key words Culture system , Dendrobium , Hybrid Cymbidium , In vitro propagation , Plant growth

regulator s , Protocorm-like body , Somatic embryo , Thin cell layer , Tissue culture

1 Introduction

Orchids are most likely the most important and interesting group

of ornamentals. Their complex ﬂ owers and pollination systems,

and the sheer size of the Orchidaceae, the largest family of the

plant kingdom, make members of this family fascinating sub-

jects to study. Recent reviews on the biotechnological aspects of

orchids show how important this group of plants in fact is [

1 , 2 ].

In conventional seed germination , orchids form a protocorm that

then develops into a plantlet [

3 ]. In vitro, in response to various

abiotic or biotic cues, orchids may also form structures, the pro-

tocorm-like body or PLB , which is the accepted de facto somatic

embryo of an orchid ( see , for example, [

4 ]). It is a structure that

resembles the true seed- and zygotic embryo -derived protocorm,

but it is derived from somatic tissue, hence the “like” term in PLB.

Nonetheless, the PLB forms a shoot and root, independent of any

surrounding tissues, and as an independent structure, making it a

372

valid case of somatic embryogenesis (SE). Thus, the term PLB

will be used throughout this chapter to synonymously represent a

somatic embryo. In addition, the initial development of a PLB

will be referred to as primary SE and the structures as primary (1°)

PLBs; then, subsequent PLBs, derived from the subculture of 1°

PLBs, will be referred to as 2° PLBs, forming within a process of

secondary SE, hereafter as 2° SE, so as to be consistent with

previous publications that deﬁ ned, and used, these terms [

5 , 6 ].

A tertiary (3°) PLB (or third- generation PLB, also derived from 2°

SE) is identical to a 2° PLB (in terms of its origin), but it is strictly

clonal, having an almost identical size, shape and dimensions, mak-

ing it the ideal explant for controlled tissue culture experiments.

Unlike 2° PLBs, which are useful starter material for establishing

repeated cycles of SE, 3° PLBs could be used in commercial

micropropagation.

Most likely the ﬁ rst orchid to have been cultured in vitro was

of the genus Cymbidium , making this an important genus to cover

within this protocol. The other genus covered by this protocol

chapter, Dendrobium , is a commercially important orchid genus

with wide-ranging medicinal properties [

7 ]. Hybrid Cymbidium

Twilight Moon ‘Day Light’ (and other cultivars) can be cultured

in vitro through three main routes, one of them being through the

use of thin cell layer s (TCLs) [

8 – 10 ]. The quantitative outcome

(i.e., PLBs) from TCLs is lower than when half-PLBs—conven-

tional explants—are used. Thus, the protocol in this chapter will

use half-PLBs. For Dendrobium , recently published protocols for

D. ‘Jayakarta’, D. ‘Gradita 31’ and D. ‘Zahra FR 62’ [

11 ], D.

‘Zahra FR 62’ [

12 , 13 ], and D. ‘Gradita 31’ [ 14 ] serve as the basis

for the protocol described herein.

2 Materials

All water used in the Cymbidium protocol is double-deionized,

ultrapure water (18 MΩ cm at 25 °C), using a Millipore

® puriﬁ er,

and is prepared fresh each time. In the Dendrobium protocols, all

water used is distilled water using a GFL Mono Water Still 2002

(Gesellschaft für Labortechnik mbH, Burgwedel, Germany) and

it is prepared fresh each time. All reagents for Cymbidium are of

tissue culture (TC) grade but the maker will differ, usually

selected on the basis of the lowest price (only three choices:

Sigma-Aldrich, St. Louis, USA; Wako Chemical Industries,

Osaka, Japan; Nacalai Tesque, Osaka, Japan). Sigma-Aldrich

products are listed below. All reagents for Dendrobium are TC

grade from Merck, Darmstadt, Germany; Sigma-Aldrich

International GmbH, St. Gallen, Switzerland; Duchefa Biochemie

B.V., Haarlem, the Netherlands.

Jaime A. Teixeira da Silva and Budi Winarto

373

The following equipment and reagents are required for the

Cymbidium protocol:

1. Petri dishes (100 mm diameter, 15 mm high) (As One, Osaka,

Japan).

2. Kinetin (Kin; Sigma-Aldrich).

3. α-Naphthaleneacetic acid (NAA; Sigma-Aldrich).

4. Tryptone (Sigma-Aldrich).

5. Bacto agar (Difco Labs, Sparks, Maryland, USA).

6. Gelrite

® (Duchefa-Biochemie).

7. Surgical blades (Hi stainless platinum or carbon steel; Feather

Safety Razor Co., Ltd., Osaka, Japan).

8. Whatman

® No. 1 ﬁ lter paper (9 cm diameter; Whatman,

Vienna, Austria).

9. Paraﬁ lm

® (SPI Supplies/Structure Probe Inc., West Chester,

PA, USA).

10. Coconut water (CW) from fresh, undamaged, green coconuts

( see Note 1 ).

11. Cool white ﬂ uorescent tubes (CWFTs): 40 W, Panasonic or

NEC, Tokyo, Japan.

The following equipment and reagents are required for the

Dendrobium protocols:

1. Vertical Pressure Steam Sterilizer Model LS-B50L-I (Huanyu

Pharmaceutical Equipment Co. Ltd., Zhangjiagang City,

Jiangzu, China).

2. Labconco Puriﬁ er™ Clean Bench (Labconco, Kansas City,

MI, USA).

3. Brand bottles (Kedaung Group Indonesia, Ungaran, Central

Java, Indonesia).

4. Erlenmeyer ﬂ asks (100 mL; Pyrex, IWAKI TE-32, Asahi Glass

License, PT. Anugerah Niaga Mandiri, Jakarta, Indonesia).

5. Petri dishes (9 cm in diameter; Normax, Rua Formigosa,

Portugal).

6. Forceps and scalpels (stainless steel; Meiden™, Tokyo, Japan).

7. Blades (BB510, Aesculap AG & Co. KG AM, Tutlingen,

Germany).

8. Mercury chloride ( HgCl

2 , Merck, Darmstadt, Germany).

9. Tween 20 (Sigma-Aldrich).

10. Thidiazuron (TDZ) (Sigma-Aldrich).

11. N

6 -benzyladenine (BA) (Sigma-Aldrich).

12. NAA (Sigma-Aldrich).

13. Gelrite

® (Duchefa-Biochemie).

2.1 Equipment

and Reagents

( Cymbidium )

2.2 Equipment

and Reagents

( Dendrobium )

Somatic Embryogenesis in Cymbidium and Dendrobium

374

14. CW (as for Cymbidium ).

15. CWFTs: SL-Shinyoku, PT, Ningbo Global Lamp, Jakarta,

Indonesia.

1. MS medium [

15 ] (Merck).

2. Growmore (32N:10P:10K, 20N:20P:20K, 6N:30P:30K; New

Century Drive, Gardena, CA, USA).

3. Rosasol medium (1.5 g/L 18N:18P:18K + 1.5 g/L

25N:10P:10K + TE) (SA Engrais, Rosier, Belgium).

4. PLB induction medium (PIM): Half-strength MS medium

containing 1.0 mg/L TDZ and 0.5 mg/L BA.

5. PLB proliferation medium 1 (PPM-1): Half-strength MS

medium containing 0.3 mg/L TDZ and 0.1 mg/L NAA

[

12 , 14 ].

6. PPM-2: Half-strength MS medium containing 0.05 mg/L

BA [

12 ].

7. PPM-3: Rosasol medium containing 150 mL/L CW [

11 ].

8. PPM-4: Growmore medium containing 100 mL/L CW [

14 ].

9. Somatic embryo proliferation medium (SEPM): Half-strength

MS medium with 0.5 mg/L TDZ and 0.5 mg/L BA ([

13 ];

Winarto et al. unpublished).

10. Shoot initiation medium (SIM): Half-strength MS medium con-

taining 1.5 mg/L TDZ, 0.5 mg/L BA, and 0.02 mg/L NAA).

3 Methods

In hybrid Cymbidium , PLBs can form spontaneously from the base

of in vitro shoot cultures that have rooted on an organically-rich

medium such as that containing banana extract. Once a single PLB

has formed, it can be extracted for the induction of new PLBs, i.e.,

neo -PLB induction [

5 ] ( see Note 2 ). In this chapter, the term neo -

PLB will not be used to avoid confusion, since neo-PLB can be a

2° PLB (formed from a 1° PLB) or 3° PLB (formed from a 2°

PLB). In Dendrobium , small shoots (±0.4 cm) that are derived

from greenhouse-grown plants will form the basal explants of the

protocol. The reader is advised to culture donor mother plants in

the greenhouse, according to the culture and environmental con-

ditions stipulated in [

12 ].

1. Excise young shoots from 3-year-old mature plants, growing

in a greenhouse, without any visible symptoms of bacterial,

fungal, or viral infection.

2. Place shoots under running tap water for 30 min. Surface ster-

ilize in 1.5 % (v/v) sodium hypochloride ( NaOCl ; 5 % active

chlorine) for 15 min. Transfer shoots to fresh sterilizing

2.3 Culture Media

3.1 General

( Cymbidium

and Dendrobium )

3.2 From

Greenhouse to In Vitro:

Sterilization and

Preparation of

Cymbidium Shoot Tips

Jaime A. Teixeira da Silva and Budi Winarto

375

solution for another 15 min. Rinse shoots off three times with

sterile distilled water (SDW; ×5 min each time).

3. In a sterile Petri dish, with a sharp sterilized surgical blade,

isolate apical meristems (0.5–1.0 mm end of terminal tips).

4. Culture on plant growth regulator ( PGR )-free half-strength

MS basal salt medium to induce shoots. 1° PLBs will form

spontaneously from the base about 1 % of rooted shoots in 4–6

months ( see Note 3 ).

5. Culture these 1° PLBs in PIM, as described next.

1. As described in [

12 ] and [ 14 ], wash axillary or apical shoots

(0.5–1.0 cm long) under running tap water for 30–60 min.

2. Immerse shoots in 1 % Tween-20 for 30 min and rinse with

sterile distilled water (SDW) ﬁ ve times (×5 min each rinse).

Surface sterilize shoots as follows: immerse in 0.05 % mercury

chloride ( HgCl

2 ) + 2–3 drops of Tween-20 for 10 min, rinse

5–6 times in SDW (×5 min each rinse). Slice off the damaged

surface of rinsed explants with a tissue culture blade (BB510,

Aesculap AG & Co. KG AM, Tutlingen, Germany).

3. Reduce shoot size by removing several leaves until ~0.4 cm

long. Culture explants in PIM. Culture dormant lateral shoot

tips (5–10 mm long) on SIM for 15–20 days. Then subculture

shoot tips in half-strength, PGR -free MS medium until healthy

shoots (approx. 0.5 cm long) form after 2–2.5 months of cul-

ture. After incubation, these shoots can be used as the explant

source for PLB initiation ( see Notes 4 and 5 ).

There are three methods to form 2° PLBs, possible by culturing ten

1° PLBs on 40 mL/100 mL ﬂ ask of PIM ( see Note 6 ). For each

method:

1. Use either Vacin and Went (VW) medium [

16 ] or Teixeira

Cymbidium (TC) medium [

17 ] supplemented with Nitsch

microelements [

18 ], 2 mg/L tryptone ; NAA and Kin are

added, each at 0.1 mg/L.

2. Add 2 % sucrose (w/w) to PIM. Adjust pH to 5.8 ± 0.1. Add

8 g/L Bacto agar .

3. Autoclave PIM at 121 °C for 21 min.

4. Incubate all cultures at 25 ± 0.5 °C in a 16-h photoperiod pro-

vided by CWFTs with a low photosynthetic photon ﬂ ux den-

sity (PPFD) of 30–40 μmol/m

2 /s ( see Note 7 ).

This method involves the natural development of clusters of 2°

PLBs using initial 1° PLB clusters without any cutting or process-

ing ( see Note 8 ).

3.3 From

Greenhouse to In Vitro:

Sterilization and

Preparation of

Dendrobium Shoot

Tips

3.4 Cymbidium : 1°

SE and Formation

of 2° PLBs from 1°

PLBs; 2° SE

and Formation of 3°

PLBs from 2° PLBs

3.4.1 Method 1

Somatic Embryogenesis in Cymbidium and Dendrobium

376

1. This method involves the use of TCLs. TCLs can be useful

when the effect of some in vitro culture factors need to be

examined on a very small explant such as a transverse TCL or

tTCL (usually 5 mm long, 5 mm wide and 0.5–1.0 mm thick)

or a longitudinal TCL or lTCL (usually 5 mm long, 1–2 mm

wide and 0.5–1.0 mm thick; occasionally an epidermal strip)

( see Note 9 ).

2. When the 1° PLB grows, 2° PLBs form on the 1° PLB, usually

at the base. Select only ideal size and uniformly shaped 2° PLBs.

3. Use a new feather blade for every 6–8 PLBs. Make a 0.5–1 mm

deep incision in the shape of a square, 3–5 × 3–5 mm in area.

Slice this area to separate the epidermal 0.5–1.0 mm in one

continuous move, thus creating an lTCL ( see Note 10 ).

4. Using a new feather blade for every 6–8 PLBs, and only using

the central 5 mm girth of the 1° PLB , make a 0.5–1 mm trans-

verse slice throughout the whole PLB, thus creating a tTCL

( see Note 10 ).

1. Method 3 is the most recommended and can be performed in

VW or TC basal medium or PIM ( see Note 11 ). 2° PLBs form

on a 45- to 60-day-old 1° PLB [

19 ], usually at the base.

2. Separated out 1° PLBs and place them in an autoclaved glass

Petri dish with a double sheet of Whatman No. 1 ﬁ lter paper

laid at the base ( see Note 12 ).

3. Slice off the top 1 mm of the 1° PLB , which contains the apical

meristem, with a feather blade. Also slice off the brown or

yellow base in contact with medium and discard it ( see Notes 12

and 13 ).

4. Slice the trimmed 1° PLB (i.e. without an apical meristem and

base) symmetrically in half to yield two half-PLBs. Place half-

PLBs cut-surface down on PIM, embedded about 1 mm into

the medium ( see Notes 14 – 17 ). After 45–60 days, 2° PLBs

form on the outer, epidermal surface of the 1° PLB. 2° PLBs

are used for 2° PLB mass production or micropropagation

( see Note 18 ).

There are two methods to form 2° and 3° PLBs.

1. To form 2° PLBs, culture small shoots on PIM in the initiation

stage. Culture 1° PLBs on any one of the two PPM (PPM-1

and PPM-2; see Note 19 ), all of which give equally successful

results. 2 % sucrose (w/w) is added to all four PPM media.

2. Adjust pH to 5.8 ± 0.1. Add 2 g/L Gelrite

® only to PIM-1.

Autoclave for 20 min at 121 °C and at 15 kPa atm. Incubate

all cultures at 24 ± 1 °C in a 12-h photoperiod provided by

CWFTs with a low PPFD of ~30 μmol/m

2 /s.

3.4.2 Method 2

3.4.3 Method 3

3.5 Dendrobium : 1°

SE and Formation

of 2° PLBs from 1°

PLBs; 2° SE

and Formation of 3°

PLBs from 2° PLBs

3.5.1 Method 1

Jaime A. Teixeira da Silva and Budi Winarto

377

3. Initiate PLBs by culturing small shoots (±0.4 cm) of D. ‘Zahra

FR 62’ and D. ‘Gradita 31’ on semisolid PIM. Periodically sub-

culture shoots every 15 days for ± 2.0 months ( see Note 20 ).

4. The method to produce 3° PLBs (Fig.

1 ) involves the natural

development of clusters of 2° PLBs using initial 1° PLB clusters

without any cutting or processing. Monthly subculture is rec-

ommended to produce vigorous PLBs ( see Notes 21 and 22 ).

1. Use PPM-2 at the initiation stage and SEPM for the prolifera-

tion stage. Add 2 % sucrose (w/w) to all media. Adjust pH to

5.8 ± 0.1. Add 2 g/L Gelrite

® . Autoclave for 20 min at 121 °C

and at 15 kPa atm.

2. Incubate all cultures are at 24 ± 1 °C in a 12-h photoperiod

provided by CWFTs with a low PPFD of ~30 μmol/m

2 /s for

initiation and multiplication of somatic embryo genic callus

(SEC) and ~10 μmol/m

2 /s for somatic embryo proliferation.

3.5.2 Method 2

Fig. 1 Method 1: Initiation and proliferation of Dendrobium ‘Zahra FR 62’ and D. ‘Gradita 31’ PLBs (i.e., somatic

embryo s). ( a ) Shoot tip as the explant source. ( b ) Shoot tip with several initial PLBs in the basal part, 3.5 months

after culture. ( c ) Initial proliferation of PLBs in half-strength MS medium containing 0.3 mg/L TDZ. The concen-

tration of TDZ is reduced to 0.1 mg/L TDZ in the third subculture. ( d ) Vigorous and green PLBs form in half-

strength MS medium with 0.3 mg/L TDZ and 0.1 mg/L NAA after the sixth subculture. ( e ) Light green PLBs in

1.6 g/L Growmore medium (32N:10P:10K) supplemented with 100 mL/L coconut water (CW) after the sixth

subculture. ( f ) PLBs harvested from half-strength MS although they can form equally well on two alternative

generic media: (1) 1.5 g/L Rosasol medium (18N:18P:18K) + 1.5 g/L Rosasol medium (25N:10P:10K + TE) con-

taining 150 mL/L CW; (2) 1.6 g/L Growmore medium (32N:10P:10K) supplemented with 100 mL/L CW ( a , c , f

reproduced from [ 12 ] with permission from Elsevier BV; b reproduced from [ 14 ] with permission from TIJSAT;

d , e are originals by Budi Winarto)

Somatic Embryogenesis in Cymbidium and Dendrobium

378

3. Initiate SEC by culturing small shoot tips (±2.0 mm in length

and 1.0–2.0 mm in diameter) on semi-solid PPM-2 medium in

9 cm Petri dishes incubated in the light for 1.5–3.0 months.

This step is genotype dependent: D. ‘Sonia Ersakul’ and

D. ‘Indonesia Raya-Ina’ = 1.5–2.0 months; D. ‘Gradita 10’ =

2.5–3.0 months (Winarto et al. unpublished data).

4. SEC derived from shoot tips is used as the explant for the next

step in the form of somatic embryo genic callus slices (SECS;

equivalent to TCLs).

5. Method 2 (Fig.

2 ) proper involves the use of SECS. Callus

forms at the base of shoots cultured in PIM-2.

6. Culture SECS (3–10 mm in diameter; 1.0–1.5 mm thick) ( see

Note 23 ) on PPM-2 and incubate for 1 month in the light.

7. Subculture SECS three times to proliferate SEC ( see Note 24 ).

8. In the fourth subculture, culture SECS on SEPM to regener-

ate 2° SEs. Cultures are placed in the light after incubation for

approx. 2.0 months ( see Note 25 ).

9. To produce a high number of 3° SEs, subculture 2–3 equally

sized and healthy 2° SEs in the same medium and incubate for

approx. 2 months in the light, but never exceed six subcultures

( see Note 26 ).

4 Notes

1. Always penetrate the coconut with a sterilized borer or drill,

pour out the CW into a sterilized container, and use fresh if pos-

sible. We never store CW at −20 °C for longer than 6 months.

2. 1° PLBs are not guaranteed to form on this medium. A ripe

banana-based medium, rich in carbohydrates and sometimes

supplemented with CW (10 %, v/v), will yield more 1° PLBs.

When shoots begin to elongate (i.e., before roots elongate),

cut off roots and transfer shoots to 0.5 % (w/v) Gelrite

® -

supplemented medium with 2 % (w/v) ripe banana and 10 %

(v/v) CW. This results in more robust plantlet growth (shoots

and roots).

3. This process/medium combination usually yields 100 % sur-

vival with this cultivar.

Fig. 2 (continued) callus resulting from the fourth subculture of SECS. ( k ) 2° somatic embryos regenerated

from two standardized 1° somatic embryos, subcultured in half-strength MS medium supplemented with

0.5 mg/L TDZ and 0.5 mg/L BA (SEPM), 1 month after culture. ( l ) Profusion of 2° somatic embryos, regenerated

from 2 to 3 standardized 1° somatic embryos subcultured on SEPM, approx. 2.0 months after culture (all

photos are originals by B. Winarto, F. Rachmawat, and N.A. Wiendi)

Jaime A. Teixeira da Silva and Budi Winarto

379

Fig. 2 Method 2: Somatic embryo genic callus slices (SECS) of several Dendrobium cultivars are used for

initiation and multiplication of SEC and somatic embryo proliferation. ( a ) Shoot tip size and position used in

initiation of somatic embryogenic callus. ( b ) Shoot tip cultured in half-strength MS medium containing 1.5 mg/L

TDZ and 0.5 mg/L BA in initial culture. ( c ) SEC regenerated in the basal part of the shoot tip, 1.5 months after

culture. ( d ) SECS, 1.0–1.5 mm thick. ( e ) A SECS initially cultured in medium indicated in ( b ). ( f ) Embryogenic

callus growth in one SECS, 10 days after culture. ( g ) Embryogenic callus regenerated from one SECS, cultured

on medium indicated in ( b ) 1 month after culture. ( h ) SECS, 1.0–1.5 mm in thickness, in the third subculture.

( i ) Embryogenic callus produced from the third subculture of one SECS, 1 month after culture. ( j ) Embryogenic

Somatic Embryogenesis in Cymbidium and Dendrobium

380

4. Initial PLB formation via callus formation or direct SE, in

terms of number, quality, and speed of initiation, is genotype

dependent.

5. SIM applied continually in semisolid or liquid medium for

approx. 2.5 months is sometimes necessary for Dendrobium

varieties that demonstrate a weak shoot initiation response.

If using this method, PLBs form more easily in the next step.

6. Always use at least 40 replicates per treatment and repeat

experiments three times for robust statistical analyses. Wherever

possible, use more than one cultivar for comparison.

7. High PPFD (>80 μmol/m

2 /s) or darkness can inhibit 2° PLB

formation. If for the experimental treatment high PPFD or

darkness are required, substitute 0.1 mg/L Kin with 1 mg/L

6-benzyladenine (BA; Sigma-Aldrich) and add 1 g/L activated

charcoal ( AC ) [

20 ]. With BA and AC, 2° PLBs form, but these

are white and not numerous; however, once transferred to

light, they regain their photosynthetic capacity. AC may mirror

a darkened natural environment of the Cymbidium in tree tops

or may absorb negative compounds, such as polyphenols,

released into the medium as a consequence of wounding [

21 ].

8. This is not a good method for testing the effect on growth of

medium factors, since it is difﬁ cult to count the number of 2°

PLBs that form per 1° PLB . In addition, these initial 1° PLB

clusters have PLBs of different sizes, developmental stages

and/or number. This method is good when one wishes to sim-

ply allow PLBs to proliferate neo -PLBs, without any experi-

mental hypothesis in mind or whenever one wishes to allow

shoots to form [

22 ].

9. This method is useful for assessing the effects of several in vitro

factors, such as ethylene inhibitors and aeration [

23 ], smoke-

saturated water [

24 ], fungal elicitors [ 25 ], jasmonates and sali-

cylic acid [

26 , 27 ], magnetic ﬁ elds [ 28 ], gelling agent and

medium additives [

29 , 30 ], and culture vessel [ 31 ] or use in

studies related to genetic engineering and transformation [

32 ,

33 ], cryopreservation and synthetic seed technology [ 34 – 37 ].

In this method, survival tends to be lower, and mortality is

higher under extreme treatments, perhaps due to smaller size,

tissue wounding, and dependence on the medium. Hence, for

propagation purposes, the TCL method is not recommended.

10. Prepare the lTCL in a single stroke. If prepared in several

strokes then the explant becomes excessively damaged and

regeneration is low. The inner tissue (sub-epidermal layers

and below) of a PLB never forms 2° PLBs [

38 ]; thus tTCLs

and lTCLs only contain epidermal tissue with 2–3 layers.

tTCLs and lTCLs dry and oxidize rapidly (within the space of

a few minutes) due to their size; thus any further damage to

Jaime A. Teixeira da Silva and Budi Winarto

381

this tissue caused during explant preparation results in rapid

browning (within a few days) and, eventually, necrosis (within

1–2 weeks) of the TCL . The feather blade should thus be

changed regularly and the cut lTCLs/tTCLs should be con-

stantly submerged in sterile, double-distilled water (SDDW).

Researchers that are new to TCL technology are advised to

spend time practicing repeatedly the preparation of such

small explants before applying them to an experimental

protocol [

39 ]. By not experimenting enough may result in

very large errors in data.

11. The basal medium (abiotic factors) [

40 ] is not as important as

the explant (biotic factors) [

41 ]. Nonetheless, the choice of

medium salts and basal medium is important [

42 , 43 ]. The use

of a half- PLB is the essentially important aspect of the method

which allows for most stable propagation of 2° PLBs.

12. The level of macro- and micronutrients, as well as the ammo-

nium/nitrate ratio, can have a profound impact on 2° PLB

production [

44 ].

13. Use one new autoclaved Petri dish for each 10–20 1° PLBs

that need to be prepared. For a total of 1000 1° PLBs, 1000 mL

of SDDW is sufﬁ cient. Pour 10–20 mL of SDDW into each

Petri dish, so that the ﬁ lter paper is always soaked with a thin

layer of SDDW and the cut surface of PLBs are always sub-

merged to avoid oxidation. The use of antioxidants in PIM can

also help [

20 ]. Never allow the PLBs to dry out (always almost

completely cover the Petri dish so that the airﬂ ow from the

clean bench does not desiccate the PLBs). Never completely

submerge the PLBs in sterile SDW as an apparent hyperhydric

response occurs to PLBs, which are extremely sensitive to

stress caused by injury, water, light, carbon source [

45 ], or

temperature. Discard any 1° PLBs that have been left standing

for more than 30 min, as an apparent hyperhydric response

occurs in SDDW.

14. In the passage from the 1°–2° PLB formation, the basal part of

the PLB is callus-like or opaque in appearance due to direct

contact with PIM. 1° PLBs should never be used for 3° PLB

production, but only 2° PLBs that form on the outer layer of

1° PLBs. Indeed, 2° PLBs are almost perfectly round, have a

more consistent shape and size, and do not have a cytologically

or morphologically distorted base.

15. Explants (1° half-PLBs) should never be placed with the intact

surface down on the medium, but simply placed on top of the

medium or slightly (0.5–1.0 mm) embedded in medium; they

should never be totally embedded into the medium, as well, as

in this case, PLBs will rarely form. This aspect needs to be

conducted uniformly ac ross experiments to avoid the distor-

tion of data.

Somatic Embryogenesis in Cymbidium and Dendrobium

382

16. Usually the “mother” PLB (i.e., the 1° PLB) will gradually die

away and turn brown (i.e., oxidize). This will take about 60–90

days to occur, depending on the cultivar. At that time, ideal

sized 2° PLBs will have formed. Following one more subcul-

tures, 2° PLBs form 3° PLBs, which can be used for experi-

mental purposes, or for micropropagation. In principle, 2°

PLBs of different sizes should never be used for experiments,

since initial PLB size strongly affects the outcome of tissue

culture experiments (Teixeira da Silva, unpublished data).

17. The sharpness of the blade is one of the most important factors

that determines the success of all three methods, especially for

the preparation of TCLs. Use sharp, feather, and robust blades

that can be autoclaved, sterilized, boiled, sterilized in 98 %

ethanol and still remain sharp for explant preparation.

18. The quantitative outcome of all three methods differs.

Quantiﬁ cation is not easy to perform with method 1, and this

method should never be used in experiments because the size,

shape and developmental stages of PLBs differ so widely in

PLB clusters. Very unfortunately, what is commonly observed

in the literature for several orchid genera is precisely the erro-

neous use of method 1 rather than methods 2 or 3. As described

in [

6 , 8 ], method 2 results in an average of about 14.5 2°

PLBs per 1° PLB lTCL and of 6 2° PLBs per 1° PLB tTCL .

The reason is the lower total surface area of tTCL than lTCL,

explained by the plant growth correction factor (PGCF) [

19 ,

46 , 47 ]. The PGCF takes into account the size of the explant,

its shape, and thus its area and thus allows hypothetical output

to be calculated based on actual data for explants of a known

size or area. In Cymbidium , two lTCLs can be prepared from

an ideal-sized 1° PLB, while five tTCLs can be prepared

from the same mother 1° PLB. Hypothetically, each sub-

culture can yield a 24,280× multiplication rate after three

consecutive subcultures (3 months each) for lTCLs. In other

words, with two initial 1° PLB lTCLs, a total of about 351,700

3° PLBs can be obtained after a 9-month period, assuming

that every single 1° and 2° PLB is used, that every single 1°

and 2° PLB survives and that every single 1° and 2° PLB is

able to differentiate. For tTCLs, these values are lower.

Hypothetically, each subculture would yield a 4620× multipli-

cation rate after three consecutive sub-cultures (3 months

each). In other words, from ﬁ ve initial 1° PLB tTCLs, a total

of about 28,100 3° PLBs can be obtained after a 9-month

period, assuming that every single 1° and 2° PLB is used, that

every single 1° and 2° PLB survives and that every single 1°

and 2° PLB is able to differentiate. Method 3 results in an

average of 8.21 2° PLBs per 1° half-PLB. Hypothetically, each

subculture can yield a 4000× multiplication rate after four

Jaime A. Teixeira da Silva and Budi Winarto

383

consecutive sub-cultures (3 months each). In other words,

with an initial two 1° half-PLBs, a total of about 36,350 3°

PLBs can be obtained after a 12-month period, assuming that

every single 1° and 2° PLB is used, that every single 1° and 2°

PLB survives and that every single 1° and 2° PLB is able to

differentiate.

19. Subculturing PLB clusters in PPM-1 and in PPM-2 allows for

the multiplication of PLBs without PLB browning [

11 , 12 ].

However, the use of PPM-3 and PPM-4 stimulates browning

in 7 % and 20 % of PLBs, respectively [

11 , 14 ].

20. A 15-day periodic subculture in the initiation stage allows

shoots to remain green and vigorous. In this state, when used

as the explant source, they easily produce SEC at the base of

shoots. After 1 month of incubation, new PLBs formed, on

average, 2.4 PLBs from one PLB in D. ‘Gradita 31’, and 2.2 in

D. ‘Zahra FR 62’ [

12 , 14 ].

21. PLB multiplication by monthly subcultures is possible for a

maximum of 8–9 subcultures, after which proliferation capacity

decreases.

22. The productivity (i.e., number of PLBs formed) of method 1

using PPM-1 and PPM-2 is higher than when PPM-3 and

PPM-4 are used. If one shoot tip (initially 0.4 mm in size)

regenerates ﬁ ve new PLBs in the initiation stage and each 2°

PLB produces ﬁ ve new PLBs (3° PLBs) in each subculture in

the multiplication stage, then 1,953,125 PLBs of D. ‘Jayakarta’

are easily produced after nine subcultures, as well as 5504 PLBs

for D. ‘Gradita 31’ and 2744 PLBs for D. ‘Zahra FR 62’.

23. The diameter of SECS in each subculture period is 3–10 mm.

In initial culture, shoot tips are about 2 mm in diameter, then

they grow up to 3 mm at the end of initiation and become

5 mm in the ﬁ rst subculture, 7 mm in the second subculture

and 10 mm in the third subculture.

24. Application of SECS can successfully produce large amounts of

SEC on PPM-2 by subculturing the SECS monthly up to three

times. SECS were used for D. ‘Sonia-Ersakul’, D. ‘Indonesia

Raya’, and D. ‘Gradita 10’ with the highest somatic embryo-

genic response (i.e., formation of SEC) and subsequent somatic

embryo formation exhibited by D. ‘Indonesia Raya’, followed

by D. ‘Sonia-Ersakul’ and D. ‘Gradita 10’ ([

13 ]; Winarto et al.

unpublished).

25. Culturing SECS on SEPM results easily in a high number of 1°

somatic embryo s (15–30/slice), derived from the fourth sub-

culture of the SECS after incubation for approx. 2.0 months

([

13 ]; Winarto et al. unpublished).

26. Production of 2° somatic embryo s (5–15/1° somatic embryo)

can be continued by culturing 2–3 uniform 1° somatic embryos

Somatic Embryogenesis in Cymbidium and Dendrobium

384

in the same medium and incubation conditions up to six

subcultures. Using this method, approximately one million

D. ‘Indonesia Raya-Ina’ and D. ‘Sonia-Ersakul’ somatic

embryos can be produced by the sixth subculture, with 10 2°

somatic embryos derived from one somatic embryo produced

in each subculture. In D. ‘Gradita 10’, 30–50 % fewer somatic

embryos are produced in the same subculture in comparison

to D. ‘Indonesia Raya-Ina’ and D. ‘Sonia-Ersakul’ ([

13 ];

Winarto et al. unpublished). Subculturing early is essential to

avoid browning, which can begin to form as early as the ﬁ rst

month after callus induction (evidenced in closely related

Pigeon orchid, Dendrobium crumenatum Swartz; [

48 ]).

Acknowledgements

The authors wish to thank Elsevier Ltd. for copyright permission

to re-use photos in Fig.

1a, c, and f from [ 12 ]. The authors also

thank Thammasat International Journal of Science and Technology

for copyright permission to reuse the photo in Fig.

1b from [ 14 ].

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Jaime A. Teixeira da Silva and Budi Winarto

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 19

Somatic Embryogenesis of Lilium from Microbulb

Transverse Thin Cell Layers

Pablo Marinangeli

Abstract

A reliable somatic embryogenesis protocol is a prerequisite for application of other plant biotechniques.

Several protocols were reported for genus Lilium , with variable success. Between them, transverse Thin

Cell Layers (tTCL) were used efﬁ ciently to induce indirect somatic embryogenesis of Lilium . Somatic

embryogenesis potential is dependent on the genotype, explant, and culture medium composition, espe-

cially as for plant growth regulators and environmental conditions. Usually, the process comprises three

phases: embryogenic callus induction, embryogenic callus proliferation and somatic embryo germination.

Somatic embryo germination can be achieved in light or dark. In the ﬁ rst case, complete plantlets are

formed, with green leaves and pseudobulb in the base. In darkness, microbulbs are formed from single

somatic embryos or clusters. A last phase of microbulb enlargement allows plantlets or microbulbs to

increase their biomass. These enlarged microbulbs do not need special acclimatization conditions when

transferred to soil and quickly produce sturdy plants. This chapter describes a protocol for somatic embryo-

genesis of Lilium using tTCL from microbulbs.

Key words Lilium , Plant tissue culture , Somatic embryo s , Transverse thin cell layer s , tTCL

1 Introduction

Lilium is a very popular cut ﬂ ower and pot plant and one of the

most important ornamental bulb worldwide. According to the

Flower Bulb Inspection Service of the Netherlands (Bloembollen-

keuringsdienst; BKD), 2.21 billion lily bulbs were produced on

3681 ha in 2010, whereas in 2014 the planted area was 3898 ha .

Other relevant lily bulb producer countries are France (401 ha),

Chile (205 ha), the USA (200 ha), Japan (189 ha), New Zealand

(110 ha), China (100 ha), and Israel (100 ha) [

1 ]. Besides its orna-

mental attributes, the increasing popularity of lilies is due to the

constant onset of novelties in terms of cultivars with superior and

distinctive features. Actually, during the last 50 years, thousands of

cultivars were developed and can be classiﬁ ed in different hybrids

388

groups. This dramatic change in the assortment of lilies was possi-

ble due to the innovative new hybrid breeding strategies that

include biotechnological tools, such as in vitro pollination and

embryo rescue, polyploidization, molecular cytogenetics and

molecular assisted breeding. Other biotechniques are not included

as breeding tools, yet. This is the case of genetic transformation,

but in the near future it is hypotesizable its key role in the incorpo-

ration of new traits to the Lilium genome [

2 ] .

Besides biotechniques directly related to breeding, there are

others assisting breeders and growers to offer an assortment of qual-

ity and quantity. These are the techniques for virus eradication,

propagation and conservation including, among others, somatic

embryo genesis. Indeed, somatic embryos are structures that allow

cloning elite material effectively in automated systems, synthetic

seed production and cryopreservation, as well as the use of them or

the embryogenic tissue as target for genetic transformation [

3 ]. So,

a reliable somatic embryogenesis (SE) protocol of Lilium is a pre-

requisite for application of other plant biotechniques. Several proto-

cols were reported for genus Lilium (Table

1 ), with variable success.

In all cases, somatic embryos were produced via indirect SE. Usually,

the process comprises three phases: embryogenic callus induction,

embryogenic callus proliferation, and somatic embryo germination

[

4 ]. SE potential is dependent on the genotype, explant, and culture

medium composition, especially as for plant growth regulator s

( PGR ), and environmental conditions [

4 , 5 ]. Most of the SE proto-

cols for Lilium makes use of the MS medium [

6 ], supplemented

with sucrose as carbon source and agar for geliﬁ cation (Table

2 ).

PGRs commonly used for inducing embryogenic calli and for prolif-

eration are auxins, as α-naphthalene acetic acid (NAA) [

4 , 5 , 7 ],

2,4- dichlorophenoxyacetic acid [

7 ], picloram [ 8 , 9 ] and dicamba [ 8 ,

9 ], and cytokinins like thidiazuron (TDZ) [ 4 ], kinetin (Kin) [ 5 ] and

N

6 -benzyladenine [ 7 , 10 , 11 ]. Somatic embryo germination can be

achieved in light or darkness. In the ﬁ rst case, complete plantlets are

formed, with green leaves and pseudobulbs at the base. In darkness,

microbulbs are formed from single somatic embryos or clusters, cul-

tured in hormone-free MS medium [

4 ]. A last phase of microbulb

enlargement in PGR-free MS medium, containing 90 g/L sucrose,

allows plantlets or microbulbs to increase their biomass. These

enlarged microbulbs do not need special acclimatization conditions

when transferred to soil and quickly produce sturdy plants [

12 ].

Thin cell layer s ( TCL ) technology consists on the in vitro cul-

ture of thin slices of tissue from different organs, and allows to

induce ﬂ owers, vegetative buds, roots, and somatic embryo s in a

very controlled pattern of organogenesis [

13 ]. About 1 mm thick

transverse slices of tissue are termed transverse TCL ( tTCL ) and

are efﬁ ciently used to induce somatic embryogenesis in Lilium

[

4 , 5 , 10 , 11 ]. This chapter describes a protocol for somatic

embryogenesis of Lilium using tTCL from microbulbs .

Pablo Marinangeli

389

Table 1

State of the art in somatic embryo genesis of Lilium

Explant Lilium species and hybrids Reference

Microbulbs, bulblets, and

bulb scales L. regale [ 7 ]

L. longiﬂ orum [

14 ]

L. formosanum [

15 ]

L. martagon [

16 ]

L. michiganense [

17 ]

L. ledebourii [

10 , 11 ]

L. davidii [

5 ]

L. longiﬂ orum [

5 ]

Longiﬂ orum x

Oriental hybrid

Asiatic hybrid

[

5 ]

Leaves L. regale [

7 ]

Seedling roots L. martagon [ 16 ]

L. x formolongi [

18 ]

L. ledebourii [

10 , 11 ]

Hypocotyls L. martagon [ 16 ]

Floral pedicels L. longiﬂ orum

Oriental hybrid [ 8 ]

Oriental hybrid [

9 ]

Styles L. longiﬂ orum

Oriental hybrid [ 8 ]

Anthers L. longiﬂ orum [

19 ]

Table 2

Composition of media used in the different steps of Lilium somatic embryo genesis by tTCL

Medium

Salts and

vitamins NAA (mg/L) TDZ (mg/L) Kin (mg/L) Sucrose (%) Agar

Bulbiﬁ cation [ 20 ] MS 0.03 – – 3 0.8

Enlargement [ 12 ] MS 0.1 – 0.1 9 0.8

Embryogenic callus

induction [

4 ] MS 1 0.2 – 3 0.8

Embryogenic callus

proliferation [

4 ] MS 1 0.1 – 3 0.8

Regeneration [

14 ] ½ MS – – – 3 0.8

MS Murashige and Skoog medium [ 6 ], NAA α-naphthaleneacetic acid, TDZ thidiazuron, Kin kinetin

Somatic Embryogenesis of Lilium

390

2 Materials

1. Different Lilium hybrids can be used for SE through this pro-

tocol. Originally it was developed for Lilium longiﬂ orum [

4 ],

but SE was achieved for other genotypes through slightly dif-

ferent protocols [

5 , 10 , 11 ]. In our laboratory, this methodol-

ogy has been successfully applied to Lilium longiﬂ orum , cv

White Heaven, Asiatic Hybrid ‘Nello’, Longiﬂ orum x Asiatic

Hybrid ‘Royal Respect’, and Longiﬂ orum x Oriental Hybrid

‘Triumphator’ ( see Note 1 ).

2. Lily bulbs can be taken from the soil after the aerial shoots die

out during autumn and winter and used directly, or they can be

harvested in autumn, stored for 1 year at −1.5 °C (after a cold

treatment during 45–60 days) and used year round.

Alternatively, bulbs can be purchased to retailers or cut ﬂ ower

growers and used directly ( see Note 2 ).

1. Laminar ﬂ ow cabinet.

2. Bunsen burner.

3. Dissecting forceps and scalpel.

4. Sterile Petri dishes 15 × 100 mm.

5. Beakers.

6. Sterile tissue paper.

7. 100–1000 mL bottles.

8. Capped test tubes (25 × 150 or 10 × 15 mm).

9. Aluminum foil.

10. Paraﬁ lm.

11. Autoclave.

12.

Stereomicroscope (if necessary, depending on the skills of the

operator).

13. pH meter.

14. Analytical balance.

15. Stirrer with hot plate.

16. Growth chamber with temperature control (25 ± 2 °C) and

light control (dark and 16-h photoperiod, at a light intensity of

40 μmol/m

2 /s provided by cool-white ﬂ uorescent tubes).

17. Greenhouse with climatic control.

1. Sterile distilled water.

2. 70 % ethanol.

3. Sodium hypochlorite or commercial bleach (e.g., Clorox

® )

with 6–8 % of active chlorine.

2.1 Plant Source

Material

2.2 Laboratory

Materials

and Equipment

2.3 Reagents,

Solutions, and

Culture Media

Pablo Marinangeli

391

4. Tween-20.

5. Culture media: Speciﬁ c media used for all the steps, from

microbulb differentiation from scale sections to shoot regen-

eration and bulb enlargement, are described in Table

2 .

3 Methods

Prepare media from the formulations in Table 2 ( see Note 1 ).

1. In an appropriate sized beaker, add distilled or deionized water

up to ½ the ﬁ nal medium volume (i.e., 500 mL for 1000 mL

medium).

2. Add mineral salts from stocks, vitamins, sucrose , and growth

regulators, stirring after each addition until the compound is

dissolved.

3. Bring to ﬁ nal volume with distilled or deionized water, mix

well, and adjust pH to 5.8 with 0.1 N NaOH or HCl.

4. Add agar , heat until gelling agent is fully dissolved, and dis-

pense into autoclavable containers. Dispense 15 mL medium

in each 25 × 150 mm tube or 5 mL medium in each 10 × 15 mm

tube. Cap tubes and place in autoclavable racks or in high-

density autoclavable polyethylene bags ( see Note 3 ).

5. Autoclave at 121 °C for 20 min (118 kPa steam pressure).

6. Store the medium in a clean area and use within 2 weeks.

1. Detach external and middle scales of healthy bulbs . Discard

external scales showing evident signs of contamination or

damage.

2. Wash carefully the scales with tap water and disinfect them by

immersion in 70 % ethanol during 1 min, followed by 20 min

in an aqueous solution of sodium hypochlorite (1.6 % active

chlorine) plus 0.1 % Tween 20.

3. Under a laminar ﬂ ow hood, rinses explants tree times with

sterile water for 2 min each, and leave them in ﬁ nal rinse water.

4. Cut scales transversally in 2–3 mm sections on a sterile tissue

paper or Petri dish. Place sections slightly embedded in the jel-

liﬁ ed bulbiﬁ cation medium, maintaining the polarity .

5. Cultivate explants in growth chamber at 25 °C in the dark dur-

ing 30–45 days until bulblets

, 3–5 mm in diameter, differenti-

ate from the base of sections.

6. Microbulbs can be used to prepare tTCL , from which to

induce SE, or they can be micropropagated in order to provide

microbulbs continuously ( see Note 4 ).

3.1 Preparation

and Sterilization

of Culture Media

3.2 Surface

Sterilization of Bulb

Scales and Culture

Somatic Embryogenesis of Lilium

392

1. In order to provide cyclic micropropagation, microbulbs

developed from scale cuttings are separated and cultivated in

enlargement medium for 45–60 days at 25 °C in the dark.

2. Enlarged microbulbs , about 5–8 mm in diameter, can be used

to obtain tTCL for SE, or used as source of microscales for

cyclic micropropagation.

1. Detach microscales from enlarged microbulbs and cultivate

them slightly embedded in bulbiﬁ cation medium.

2. Cultivate in growth chamber at 25 °C in darkness during

30–45 days until bulblets , 3–5 mm in diameter, differentiate

from the base of microscales .

3. Microbulbs can be used to obtain tTCL to induce SE or to

continue cyclic micropropagation.

1. Remove microbulbs from tubes or Petri dishes. Cut roots and

etiolated leaves, while microscales should remain.

2. Excise 0.8–1.0 mm thick tTCL from the base of microbulbs

and place them with the inverted polarity on embryogenic cal-

lus induction medium.

3. Cultivate in growth chamber at 25 °C in the dark with 30-day

subcultures.

4. Remove embryogenic callus from the explants and cultivate them

in embryogenic callus proliferation medium, in growth chamber

at 25 °C in the dark with 30-day subcultures ( see Note 5 ).

1. Transfer proliferating embryogenic callus to regeneration

medium.

2. Cultivate in growth chamber at 25 °C with a 16-h photoperiod

at a light intensity of 40 μmol/m

2 /s.

3. Transfer to fresh medium every 30 days until microshoot

development.

1. Transfer microshoots to enlargement medium and cultivate

them in a growth chamber at 25 °C in darkness.

2. Cultivate during 60 days with one subculture.

3. Remove microbulbs from culture containers, wash under tap

water to remove medium and plant directly in soil or substrate

( see Note 6 ).

4 Notes

1. SE depends on the genetic background of the donor plant. The

response of different species and cultivars of Lilium to embryo-

genic callus induction and proliferation is variable and it is pos-

3.3 Enlargement

of Microbulbs

3.4 Bulbiﬁ cation

3.5 Embryogenic

Callus Induction

and Proliferation

3.6 Germination

of Somatic Embryos

and Plant

Regeneration

3.7 Bulbiﬁ cation

and Soil Transfer

Pablo Marinangeli

393

sible that would be necessary an adjustment of the culture

conditions. The main factors affecting SE are the type and con-

centration of PGR . So, it is recommended an adjustment of the

concentrations of NAA (0.1–1 mg/L) and TDZ (0.1 –0.4 mg/L)

when working with novel material [

4 , 5 , 10 , 11 ].

2. When lily bulbs are damaged or dehydrated, high frequency of

contamination appears during in vitro culture because disinfec-

tion is not efﬁ cient, reaching even 100 % of loss. In this situa-

tion, it is possible to carry out an ex vitro propagation of bulbs

through scaling technique [

13 ], during which a strong disin-

fection of scales is done with disinfectants, fungicides, and

acaricides. Furthermore, the production of new healthy organs

allows obtaining a material suitable for establishment in vitro,

the explants being the microscales from bulblets differentiated

at the base of the scales.

3. During the introduction in vitro it is absolutely necessary to

use single culture tubes because the contamination is usually

high. During the step of embryogenic callus proliferation and

the embryos germination , it is possible to use Petri dishes to

cultivate explants, due to the possibility to save space and cul-

ture medium. This requires preparing and sterilizing the cul-

ture medium in ﬂ asks with plastic cap, and then pouring it into

sterile disposable Petri dishes while still melted.

4. Microbulbs , differentiated from scale sections of the original

bulb , can be used for SE, but if year-round work is necessary, a

continuous source of microbulbs is necessary. So, it is recom-

mended following the cyclic micropropagation of Lilium in

the dark, as mentioned in Subheadings 3.2, 3.3, and 3.4.

5. Embryogenic callus proliferation can be done both in solid and

in liquid medium by cultivating embryogenic calli in either

agar -solidiﬁ ed or liquid MS media, containing 1.0 mg/L NAA

and 0.2 mg/L TDZ. However, the number of somatic embryo s

derived from embryogenic calli cultured in liquid medium

often shows to be more than in solidiﬁ ed medium [

14 ].

6. In some cultivars, microbulbs develop dormancy during

enlargement. In this case, break of dormancy is possible by

storing microbulbs at 4–7 °C from 45 days ( L. longiﬂ orum ,

Longiﬂ orum x Asiatic, and Asiatic hybrids) to 60 days

(‘Oriental’ and ‘Oriental x Trumpet’ hybrids). Dormancy

release can be done in the same culture container or in humid

peat moss within plastic bags, or in plastic containers covered

with ﬁ lm.

Somatic Embryogenesis of Lilium

394

References

1. Grassotti A, Gimelli F (2011) Bulb and cut

ﬂ ower production in the genus Lilium : current

status and the future. Acta Hort 900:21–35

2. Van Tuyl JM, Arens P (2011) Lilium : breeding

history of the modern cultivar assortment. Acta

Hort 900:223–230

3. Marinangeli P (2012) Micropropagazione delle

geoﬁ te ornamentali. Italus Hortus 19:51–56

4. Nhut DT, Van Le B, Minh NT, Teixeira da Silva

J, Fukai S, Tanaka M, Van Tran Thanh K (2002)

Somatic embryogenesis through pseudo-bulb-

let transverse thin cell layer of Lilium longiﬂ o-

rum . Plant Growth Regul 37:193–198

5. Yin ZF, Bi WL, Chen L, Wang QC (2014)

Plant regeneration and cryopreservation of

Lilium . Acta Hort 1027:185–196

6. Murashige T, Skoog F (1962) A revised medium

for rapid growth and bio assays with tobacco tis-

sue cultures. Physiol Plant 15:473–497

7. Pelkonen VP, Kauppi A (1999) The effect of

light and auxins on the regeneration of lily

( Lilium regale Wil.) cells by somatic embryo-

genesis and organogenesis. Int J Plant Sci

160:483–490

8. Tribulato A, Remotti PC, Lofﬂ er HJM, van

Tuyl JM (1997) Somatic embryogenesis and

plant regeneration in Lilium longiﬂ orum

Thunb. Plant Cell Rep 17:113–118

9. Tribulato A, Branca F (2011) Somatic embryo-

genesis from ﬂ ower pedicels of oriental lilies.

Acta Hort 900:369–376

10. Bakhshaie M, Babalar M, Mirmasoumi M,

Khalighi A (2010) Effects of light, sucrose, and

cytokinins on somatic embryogenesis in Lilium

ledebourii (Baker) Bioss. via transverse thin

cell-layer cultures of bulblet microscales. J Hort

Sci Biotechnol 85:461–464

11.

Bakhshaie M, Babalar M, Mirmasoumi M,

Khalighi A (2010) Somatic embryogenesis and

plant regeneration of Lilium ledebourii (Baker)

Boiss., an endangered species. Plant Cell Tiss

Org Cult 102:229–235

12. Marinangeli PA, Curvetto NR (1997)

Increased sucrose and salt concentrations in the

culture medium improve growth of microprop-

agated Lilium bulblets. Biocell 21:161–164

13. Marinangeli PA, Hernández LF, Curvetto NR

(2003) Bulblet differentiation from Lilium lon-

giﬂ orum Thunb. bulb-scales. J Am Soc Hort

Sci 128:324–329

14. Nhut DT, Hanh NTM, Tuan PQ, Nguyet

LTM, Tram NTH, Chinh NC, Nguyen NH,

Vinh DN (2006) Liquid culture as a positive

condition to induce and enhance quality and

quantity of somatic embryogenesis of Lilium

longiﬂ orum . Sci Hortic 110:93–97

15. Nakano M, Sakakibara T, Suzuki S, Saito H

(2000) Decrease in the regeneration potential

of long-term cell suspension cultures of Lilium

formosanum Wallace and its restoration by the

auxin transport inhibitor, 2, 3, 5- triiodobenzoic

acid. Plant Sci 158:129–137

16. Kedra M, Bach A (2005) Morphogenesis of

Lilium martagon L. explants in callus culture.

Acta Biol Cracov Bot 47:65–73

17. Ault JR, Siqueira SS (2008) Morphogenetic

response of Lilium michiganense to four auxin-

type plant growth regulators in vitro.

HortScience 43:1922–1924

18. Ho CW, Jian WT, Lai HC (2006) Plant regen-

eration via somatic embryogenesis from sus-

pension cell cultures of Lilium × formolongi

Hort. using a bioreactor system. In Vitro Cell

Dev Biol Plant 42:240–246

19. Arzate-Fernandez AM, Nakazaki T, Okumoto

Y, Tanisaka T (1997) Efﬁ cient callus induction

and plant regeneration from ﬁ laments with

anther in lily ( Lilium longiﬂ orum Thunb.).

Plant Cell Rep 16:836–840

20.

Curvetto NR, Marinangeli PA, Mockel G

(2006) Hydrogen peroxide in the micropropa-

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with a traditional methodology. Biocell

30:497–500

Pablo Marinangeli

395

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 20

Somatic Embryogenesis and Plant Regeneration

of Brachiaria brizantha

Glaucia B. Cabral , Vera T. C. Carneiro , Diva M. A. Dusi ,

and Adriana P. Martinelli

Abstract

The genus Brachiaria (Trin.) Griseb. belongs to the family Poaceae, order Poales, class Monocotyledonae.

In Brachiaria brizantha (Hochst. ex A. Rich.) Stapf., embryogenic callus can be induced from seeds from

apomictic plants, which results in high frequency somatic embryo development and plant regeneration. We

report here a detailed protocol for callus induction from apomictic seed; followed by in vitro morphogen-

esis (somatic embryo and bud differentiation), plant regeneration, and acclimatization in the greenhouse.

Important details regarding the positioning of seeds for callus induction and precautions to avoid endo-

phytic contamination and the occurrence of albino plants are presented.

Key words Albino plants , Apomixis , Caryopses , Endophyte contamination , Forage grass , In vitro

culture , Monocot

1 Introduction

The Brachiaria genus belongs to the family Poaceae, order Poales

of the class Monocotyledonae. This genus shows a hundred species

of grasses [

1 ], and is cultivated as forage in tropical and subtropical

countries. Brachiaria brizantha cv. Marandu is a key forage in beef

cattle production in Brazil, the world’s largest beef exporter. B.

brizantha shows aposporic apomixis , an asexual mode of reproduc-

tion by seeds. The apomictic plants are thus clones of the mother

plant, reducing the possibility of their use in classical breeding [

2 ].

Furthermore, similarly to many apomicts [

3 ], apomixis is related to

polyploidy, with sexual plants being diploid and apomictic plants

tetraploids. These characteristics hinder breeding of apomictic

plants [

4 ].

Alternatives to conventional breeding of B. brizantha would

involve the introduction of genes of interest by genetic transforma-

tion, which relies on the availability of in vitro plant regeneration

protocols. In monocots , in vitro plant regeneration has been

396

primarily achieved through somatic embryo genesis [ 5 , 6 ]. Evidences

of different morphogenic responses from the same explant, under

induction by different concentrations and ratios of auxin:cytokinin,

have been reported in species of the Poaceae family such as sor-

ghum ( Sorghum bicolor (L.) Moench.), minor millet ( Paspalum

scrobiculatum L.), sugarcane ( Saccharum ofﬁ cinarum L.), and baby

bamboo ( Pogonatherum paniceum Lam. Hack.) [

7 ]. The occur-

rence of monopolar and bipolar embryos in sugarcane cultures,

which has been earlier described as two pathways [

8 ], was later

deﬁ ned as organogenesis and somatic embryogenesis [

9 ]. In B. bri-

zantha cv. Marandu, high in vitro morphogenetic efﬁ ciency was

observed from seeds, with 73 % of isolated apomictic embryos

forming embryogenic cultures, and 67 % of the calli regenerating

plants [

10 ]. The anatomy of somatic embryos, induced from in vitro

cultivated seeds, conﬁ rmed this morphogenetic route [

11 ]. Multiple

shoot formation was also reported in basal segments obtained from

micropropagated plantlets of cv. Marandu [

12 ], a system that was

used for Brachiaria plant recovery after colchicine treatment for in

vitro chromosome duplication. Somatic embryo genesis and organ-

ogenesis in B. brizantha is inﬂ uenced by several factors such as

genotype, explant type, and culture conditions [

13 ].

In this chapter, an efﬁ cient protocol of somatic embryo genesis

induction from apomictic mature seeds is reported.

2 Materials

1. Mature seeds of Brachiaria brizantha cv. Marandu.

2. 70 % ethanol solution (v/v) in water.

3. 2.5 % sodium hypochlorite ( NaOCl ) solution (v/v) in water.

4. Tween 20™.

5. Distilled water, sterilized by autoclaving at 121 °C for 20 min.

6. Conic plastic tubes, sterile, 50 mL.

7. Laminar ﬂ ow hood.

8. Filter paper placed in Petri dishes, sterilized by autoclaving at

121 °C for 20 min.

9. Petri dishes, 100 × 20 mm.

10. Scalpel blades, scalpel handles, and tissue forceps.

11. Plastic ﬁ lm, Paraﬁ lm

® M type.

12. Stock solution of 2,4-dichlorophenoxyacetic acid (2,4-D);

benzylaminopurine (BAP); kinetin (KIN); naphthaleneacetic

acid (NAA), and gibberellic acid ( GA

3 ), each one at 1 mg/mL.

13. Vessels for plant tissue culture (e.g., Magenta™ or babyfood jars).

2.1 Plant Material

and Equipment

Glaucia B. Cabral et al.

397

14. Incubator or growth chamber with controlled temperature

and photoperiod.

15. Mixture of sand:soil:vermiculite (1:1:1, v/v), sterilized by

autoclaving at 121 °C for 40 min.

16. Soil fertilized with superphosphate and organic matter.

1. Induction medium ( M1.3 ): Murashige and Skoog (MS) basal

medium [

14 ], 3 % sucrose , 300 mg/L casein hydrolysate , 3

mg/L 2,4-D, 0.2 mg/L BAP, 14 g/L agar , pH 4.2 ( see Note

1 ), poured in Petri dishes [

13 ].

2. Differentiation medium ( DM ): MS basal medium with ½

strength of major salts, 2 % sucrose , 300 mg/L casein hydroly-

sate , 0.5 mg/L 2,4-D, 0.05 mg/L BAP, 14 g/L agar , pH 4.2

( see Note 1 ), poured in Petri dishes.

3. Regeneration medium ( MS3 ): MS basal medium with 3 %

sucrose , 300 mg/L casein hydrolysate , 0.5 mg/L NAA, 1

mg/L BAP, 2.5 mg/L KIN, 14 g/L agar , pH 4.2 ( see Note

1 ), poured in Petri dishes [

13 ].

4. Elongation and rooting medium ( MMP ): MS basal medium

with ½ strength major salts, 2 % sucrose , 100 mg/L casein

hydrolysate , 0.5 mg/L KIN, 0.2 mg/L NAA, 0.2 mg/L GA

3 ,

0.7 % agar , pH 5.8, poured in Magenta™ boxes (30 mL in

each box), or other vessels for plant tissue culture [

13 ].

The pH of media is adjusted to 5.8 with 1 N KOH or to 4.2

with 1 N HCl prior to autoclaving. Medium is autoclaved at

121 °C, for 20 min. GA

3 is ﬁ lter sterilized and added to media

after autoclaving.

3 Methods

1. The seeds of B. brizantha should be dehusked manually with

the aid of a forceps (Fig.

1a ) or a seed stripper. Select the well-

formed and unblemished seeds with the aid of a stereomicro-

scope to avoid contamination. Use around 300 seeds per

treatment ( see Note 2 ).

2. Decontamination of the selected and dehusked seeds (Fig.

1b )

should be carried out in a laminar ﬂ ow hood, using a previ-

ously autoclaved beaker, with 100 mL of 70 % ethanol for 5

min, followed by 100 mL of a 2.5 % sodium hypochlorite solu-

tion with two drops of Tween 20 for 30 min; stir the solution

with seeds repeated times.

3. Rinse the seeds thoroughly, six times, with distilled autoclaved

water.

4. Dry the seeds on autoclaved ﬁ lter paper.

2.2 Culture Media

for Brachiaria

brizantha Somatic

Embryogenesis

via Callus Formation

Somatic Embryogenesis of Brachiaria Brizantha

398

5. Inoculate the seeds onto induction medium (M1.3) poured in

Petri dishes (100 × 20 mm), 10–12 seeds per plate. To avoid

condensation, leave the Petri dishes open in the hood for

15 min to remove excess of moisture. Seal the plates with

Paraﬁ lm

® and incubate them upside down, in the dark, at

25 ± 2 °C for 3–4 weeks ( see Note 3 ).

6. After 5 days, mature seeds cultured in M1.3 induction medium

present a swollen scutellum and embryo axis (Fig.

1c ). The

swelling is followed by proliferation of friable callus, after 2

weeks of induction, on the upper surface of the scutellum, and

an opaque white structure is generally observed (Fig.

1d ),

Fig. 1 Selection of Brachiaria brizantha cv. Marandu mature seeds and somatic embryo genesis induction: ( a )

selection of mature seeds for manually dehusking; ( b ) peeled mature seeds showing in one side the apomictic

embryo ( arrow head ) and in the opposite side the hilum ( arrow ); ( c ) 5-day-old mature seed cultivated in callus

induction medium, showing the swollen scutellum ( asterisk ) and embryo axis ( arrow ) of the apomictic seed

embryo; ( d ) friable callus grown on the scutellum surface of the seed ( arrow ); ( e ) multiplication of embryo-

genic callus. Bars: a = 6 mm, b = 3 mm, c = 1 mm, d = 2 mm, e = 1 mm

Glaucia B. Cabral et al.

399

which is the scutellum of a somatic embryo which, in turn,

repetitively produces new scutelli if the calli are subcultured to

induction medium M1.3 (Fig.

1e ) ( see Note 4 ).

7. To obtain well-differentiated somatic embryo s, transfer the

3- to 4-week-old induced calli from induction medium (M13)

to differentiation medium (DM), following step 7 ; however,

to obtain plant regeneration skip step 7 and go directly to

step 8 . Seal the Petri dishes with Paraﬁ lm

® and incubate them

upside down, in the dark, at 27 ± 2 °C for 3–4 weeks. In this

step visible differentiation of the somatic embryos is observed.

Add this step for a better visualization of somatic embryo

differentiation.

8. To obtain plant regeneration, transfer the 3- to 4-week-old

induced calli from induction medium (M1.3) to regeneration

medium (MS3), spreading them well on the medium, approxi-

mately 8–10 calli per Petri dish ( see Note 5 ).

9. Incubate the Petri dishes upside down in a culture room or

incubator, at 35 μmol/m

2 /s, 14 h photoperiod and 27 ± 2 °C

for 3 days, then move the plates to a higher light intensity con-

dition (70 μmol/m

2 /s) for 4 weeks ( see Note 6 ).

10. Germinating somatic embryo s, 2–3 cm long, with developing

leaves, are transferred to elongation and rooting medium

(MMP), in Magenta™ boxes (30 mL in each box) and kept in

a culture room at 25 ± 2 °C at 70 μmol/m

2 /s and 14 h photo-

period, for 3–4 weeks.

11. For acclimatization, in vitro-rooted shoots are carefully washed

to remove the agar , and then transferred ﬁ rst to plastic pots,

containing vermiculite, and covered with a plastic bag to main-

tain high humidity (4–5 days), and afterwards to pots contain-

ing a mixture of sand:soil:vermiculite (1:1:1, v/v) in the

greenhouse with natural light and temperature ( see Note 7 ).

4 Notes

1. Endophytes are very common in Brachiaria spp. The acidic

pH of the induction and regeneration medium (4.2) favors the

reduction of endophytic bacteria contamination, enabling lon-

ger term B. brizantha in vitro culture, compared to culture at

pH 5.8. However, at pH 4.2 agar solidiﬁ cation can be difﬁ cult

and, to avoid this problem, we suggest the use of type A agar

at twice the usual concentration (1.4 %).

2. Selection of well-formed and non-damaged seeds with the aid

of a stereomicroscope is very important to prevent contamina-

tion, considering that B. brizantha seeds show a high unviable

rate (30 %), and the viable seeds usually present fungi in the

Somatic Embryogenesis of Brachiaria Brizantha

400

endosperm if the storage conditions are not adequate. Seeds

should be stored under refrigeration, in a desiccator.

3. Use 100 × 20 mm Petri dishes for better aeration. It is highly

recommended to prepare and pour M1.3 medium in Petri

dishes a day or two before inoculating the seeds, to reduce

condensation and contamination. Condensation in plates

should also be avoided when sowing the seeds. If there is any

condensation, open the Petri dishes with medium in the hood

and leave them open for 15 min. Brachiaria seeds should be

positioned preferably with the embryo axis side up, i.e., the

hilum side, which is visible (Fig.

1b ), should be in contact with

the culture medium. Place the seeds, one by one, applying a

slight pressure in the culture medium without submerging

them, so that they do not detach from the medium due to the

growth of callus and/or germination of the embryo. If the

seed is not in a close contact with the medium, instead of form-

ing callus, the embryo germinates or root formation is

observed; thus, the seed needs to be slightly pressed into the

medium. The quality of the primary callus depends entirely on

the seed quality and the induction process.

4. M1.3 medium produces a high percentage of embryogenic

calli in a 2,4-D concentration ranging from 2 to 4 mg/L. We

suggest avoiding a long-term maintenance of Brachiaria bri-

zantha embryogenic calli in the presence of 2,4-D, due to a

high probability of subsequent formation of 100 % albino

plants in 4-month-old embryogenic calli [

13 ] (Fig. 2a, b ).

Moreover, recently, it was shown that, in ruzigrass ( Brachiaria

ruziziensis ), 4-month-old embryogenic callus generated poly-

ploids, while all regenerants derived from 2-month-old

embryogenic calli were diploid, suggesting that 2,4-D pro-

moted not only the formation of somatic embryo genesis, but

also duplication of chromosomes at early stages of embryo-

genic callus formation [

15 ]. These outcomes indicate that

2-month-old or younger embryogenic calli are best suited for

Brachiaria spp. When the supplementation of auxin decreases,

there is a rapid differentiation into embryos and different pat-

terns of distribution in the same explant may occur due to local

accumulations of auxin. Auxin transport and accumulation

may also have an inﬂ uence in somatic embryo differentiation in

Brachiaria . Therefore, to obtain well differentiated somatic

embryos, we highly suggest transferring the induced calli to

differentiation medium (DM), and after 3–4 weeks, embryos

have a cream-colored embryo axis with coleoptile surrounding

the shoot apical meristem of the somatic embryos, each cole-

optile containing one shoot meristem (Fig.

2c ). The embryo

proper or embryo axis is enveloped with an opaque white-col-

ored, isolated, well- differentiated scutellum, and in some cases

show fused scutelli (Fig.

2c ).

Glaucia B. Cabral et al.

401

5. In MS3 medium plant regeneration is obtained from up to 54

% of the seeds and around 90 % of induced calli. Detailed

observations of calli show that two regeneration patterns are

observed after transferring the calli to regeneration medium:

(1) complete plantlets originating from isolated somatic

embryo s (Fig.

2d ), and (2) multiple buds formed from the api-

cal meristem of somatic embryos in the presence of cytokinins

in the MS3 regeneration medium (Fig.

2e, f ), producing mul-

tiple shoots (Fig.

2g ). The purple pigmentation observed in

buds and shoots indicates a stress-induced anthocyanin pro-

duction in the leaf tips of B. brizantha cultivated in vitro under

light conditions. For reducing this stress the explant should

initially be cultured for 3 days at a reduced light intensity,

around 30 μmol/m

2 /s, returning to a higher light intensity

(70 μmol/m

2 /s), for plantlet development.

6. If two light intensity conditions are not available, as an alterna-

tive pile up the Petri dishes at 70 μmol/m

2 /s

1 for the initial

period of 3 days at reduced light intensity, then spread the

plates side by side at the same light intensity of the climatic

chamber. The initial lower light intensity helps to prevent

anthocyanin accumulation.

7. Regenerated plantlets from this protocol show a morphologi-

cal pattern of growth, ﬂ owering, and seed production, similar

to naturally propagated plants (Fig.

2h ).

Fig. 2 Plant regeneration of Brachiaria brizantha cv. Marandu: ( a ) albino shoots regenerating from embryogenic

callus; ( b ) elongating albino shoots; ( c ) differentiated scutellum and embryo axis of somatic embryo ( arrow );

( d ) plantlet originated from an isolated somatic embryo; ( e ) multiple buds formed in an embryogenic callus

( arrow ); ( f ) multiple buds; ( g ) multiple shoots elongating from clumps of buds; ( h ) tiller plant ( left ) and in vitro

plant after acclimatization ( right ). Bars: a = 1 mm, c = 0.5 mm, d = 2 mm, e = 2 mm, f = 1 mm

Somatic Embryogenesis of Brachiaria Brizantha

402

Acknowledgements

G.B.C. thanks Embrapa for a doctoral fellowship. A.P.M. acknowl-

edges CNPq for a research fellowship (310.612/2011-0).

References

1. Do Valle CB, Savidan Y (1996) Genetics, cyto-

genetic and reproductive biology of Brachiaria.

In: Miles JW, Maass BL, Do Valle CB (eds)

Brachiaria: biology, agronomy and improve-

ment, vol 1. CIAT Campo. Grande-Brasil

Embrapa, Cali-Colombia, pp 147–163, 288

2. Lutts S, Ndikumana J, Louant B (1994) Male

and female sporogenesis and gametogenesis in

apomictic Brachiaria brizantha , Brachiaria

decumbens and F1 hybrids with sexual colchi-

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Euphytica 78:19–25

3. Asker SE, Jerling L (1992) Apomixis in plants.

CRC Press, Boca-Raton, 298p

4. Do Valle CB, Bonato ALV, Pagliarini MS,

Resende RMS, Jank L (2004) Apomixia e sua

utilização no melhoramento de Brachiaria . In:

Carneiro VTC, Dusi DMA (eds) Clonagem de

plantas por sementes: estratégias de estudo da

apomixia. Embrapa Recursos Genéticos e

Biotecnologia, Brasília, pp 47–65, 126p

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cereal protoplasts. Theor Appl Genet 56:97–99

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the challenge, the debate, and the solution – a

historical perspective. In Vitro Cell Dev Biol

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7. Wang W, Zhao X, Zhuang G, Wang S, Chen F

(2008) Simple hormonal regulation of somatic

embryogenesis and/or shoot organogenesis in

caryopsis cultures of Pogonatherum paniceum

(Poaceae). Plant Cell Tiss Org Cult 95:57–67

8. Burrieza HP, Lopez-Fernandez MP, Chiquieri

TB, Silveira V, Maldonado S (2012)

Accumulation pattern of dehydrins during

sugarcane (var. SP80.3280) somatic embryo-

genesis. Plant Cell Rep 31:2139–2149

9. Falco MC, Mendes BMJ, Tulmann Neto A,

Appezatto-da-Glória B (1996) Histological

characterization of in vitro regeneration of

Saccharum sp. R Bras Fisiol Veg 8:93–97

10. Silveira ED, Rodrigues JCM, Cabral GB, Leite

JA, Costa SS, Carneiro VTC (2003) Evaluation

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brizantha transformation. J Plant Biotechnol

5:87–93

11.

Lenis-Manzano SJ, Araujo ACG, Valle CB,

Santana EF, Carneiro VTC (2010) Histologia

da embriogênese somática induzida em embriões

de sementes maduras de Urochloa brizantha

apomítica. Pesq Agropec Bras 45:435–441

12. Pinheiro AA, Pozzobon MT, Do Valle CB,

Penteado MIO, Carneiro VTC (2000)

Duplication of the chromosome number of

diploid Brachiaria brizantha plants using col-

chicine. Plant Cell Rep 19:274–278

13. Cabral GB, Carneiro VTC, Lacerda AL, Do

Valle CB, Martinelli AP, Dusi DMA (2011)

Somatic embryogenesis and organogenesis in

apomictic and sexual Brachiaria brizantha .

Plant Cell Tiss Org Cult 107:271–282

14. Murashige T, Skoog F (1962) A revised medium

for rapid growth and bioassays with tobacco tis-

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15. Ishigaki G, Gondo T, Rahman MM, Umami N,

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Everard). J Jpn Grassl Sci 60:24–30

Glaucia B. Cabral et al.

Part III

Protocols of Somatic Embryogenesis

in Selected Forest Trees

405

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 21

Somatic Embryogenesis in Pinus spp.

Itziar Aurora Montalbán , Olatz García-Mendiguren ,

and Paloma Moncaleán

Abstract

Somatic embryogenesis (SE) has been the most important development for plant tissue culture, not only

for mass propagation but also for enabling the implementation of biotechnological tools that can be used

to increase the productivity and wood quality of plantation forestry. Development of SE in forest trees

started in 1985 and nowadays many studies are focused on the optimization of conifer SE system. However,

these advances for many Pinus spp. are not sufﬁ ciently reﬁ ned to be implemented commercially. In this

chapter, a summary of the main systems used to achieve SE in Pinus spp. is reported.

Key words Conifer , Forest biotechnology , In vitro , Plant tissue culture , Propagation , Somatic

embryo s

1 Introduction

During the last years, one of the main objectives of conifer genetic

improvement programs has been the development and application

of biotechnological tools, able to achieve production systems of

elite plants adapted to different environmental conditions in the

future scenery of climate change. In vitro propagation systems

have been one of the most studied aspects worldwide in programs

of genetic improvement. In this sense, although the utility of

in vitro organogenesis using juvenile material has been developed

and optimized in some Pinus species [

1 – 5 ], the high cost of the

process is still a limitation for mass production on a commercial

scale. Other systems to achieve in vitro propagation of Pinus spp.

adult trees have been developed [

6 – 10 ], but changes in the attri-

butes of resulting plants have sometimes been observed and reju-

venation of the material has been transitory in in vitro conditions.

For the abovementioned reasons, propagation via somatic embryo-

genesis (SE) has many advantages:

406

– It captures all speciﬁ c gene combinations of selected individu-

als, giving high genetic gain [

11 ];

– Combined with cryopreservation of the embryonal masses

( EM ) and the selection of elite clones in ﬁ eld tests, it enables

implementing multivarietal forestry [

11 ];

– It is an ideal system for genetic transformation, through initia-

tion of somatic embryo s from single cells [

12 ];

– It offers the capability to produce unlimited numbers of plant-

lets from somatic embryo s [

13 , 14 ] and artiﬁ cial seed s [ 15 ].

Although EM initiation protocols are now fairly well estab-

lished, maturation of EM into cotyledonary normal somatic

embryo s is not always successful in Pinus spp. Problems such as

low or asynchronous embryo production [

16 , 17 ], abnormal mor-

phology, or poor root development have also been reported for P.

pinea [

18 ] and P. kesiya [ 19 ]. In conifer SE, physical and chemical

conditions are factors that should be studied carefully. In this sense,

culture medium composition takes on special importance in SE,

being a substitute of the megagametophyte , supplying adequate

amounts of nitrogen and carbon [

20 ]. For this reason, studies were

carried out to analyse the effect of culture conditions in SE of

Pinus spp. [

21 – 24 ]. Thanks to these studies, a considerable increase

of the number and quality of the produced Pinus somatic plantlets

has been achieved [

23 ].

In this chapter, recent studies focused on the development and

optimization of successful protocols of SE in various Pinus species

are described.

2 Materials

One-year-old green female cones, enclosing immature zygotic

embryo s of Pinus spp. at the precotyledonary stage [

21 ], are col-

lected from open or control pollinated trees ( see Note 1 ). The

cones are stored at 4 °C until processing ( see Note 2 ). Cones are

usually processed within a week, although they can be stored for

more than one month with no detriment in SE initiation rates

[

13 ]. Recently, it has also been possible to initiate EM from dif-

ferentiated cells in epicotyledonary region of post-cotyledonary

zygotic embryos [

25 ].

1. Initiation, proliferation, and maturation phases of SE are usu-

ally carried out in the same basal medium. As for macroele-

ments, microelements and vitamins, different media

formulation and their modiﬁ cations are used depending on the

species [

24 , 28 ], i.e.:

2.1 Plant Material

2.2 Media

Itziar Aurora Montalbán et al.

407

– DCR [ 26 ] with P. nigra [ 27 ], P. sylvestris [ 28 ], P. palustris

[

29 ], P. patula [ 30 ], and P. brutia [ 16 ];

– EDM [

31 ], LP [ 32 ] and Glitz [ 33 ] for P. radiata [ 15 ,

23 , 33 ];

– 505 [

34 ] for P. taeda [ 35 ];

– LV [

36 ] for P. pinaster [ 37 ], P. monticola [ 38 ], P. pinea

[

18 ], P. strobus [ 39 ], and the hybrid P. rigida x P. taeda

[

40 ]. Moreover, different modiﬁ cations of culture media

described are used, depending on the species [

41 ].

2. As a carbon source, sucrose in P. halepensis [

22 ], P. pinaster

[

37 ], P. pinea [ 18 ] and P. strobus [ 39 ], and/or maltose in P.

densiﬂ ora [

42 ], P. kesiya [ 43 ], P. patula [ 30 ] and P. taeda [ 34 ]

are used in concentrations ranging: (i) from 10 g/L in P.

armandii [

44 ] and P. luchuensis [ 45 ] to 30 g/L in P. bun-

geana [

12 ], P. pinaster [ 37 ], and P. radiata [ 21 , 33 ] for initia-

tion and proliferation, and (ii) from 30 g/L in P. monticola, [

38 ]

and P. luchuensis [

45 ] to 60 g/L in P. pinea [ 18 ], P. radiata

[

23 ], and P. strobus [ 39 ] for maturation ( see Note 3 ).

3. Gellan gum ( Gelrite

® or Phytagel

® ) is added to the medium.

The concentration of gellan gum varies, depending on the spe-

ciﬁ c phase of SE process from 2 g/L in P. brutia [

16 ] to 4 g/L

in P. pinea [

18 ] and P. monticola [ 38 ] for the initiation stage,

from 3 g/L in P. oocarpa [

46 ] to 5.5 g/L in P. radiata [ 21 ] for

the proliferation stage, and from 4 g/L in P. nigra [

47 ] to 10

g/L in P. pinaster [

37 ], P. rigida x P. taeda [ 40 ], and P. sylves-

tris [

48 ] for the maturation stage ( see Note 3 ). Moreover,

polyethylene glycol ( PEG ) is used in the maturation stage of

species such as P. armandii [

44 ], P. brutia [ 16 ], P. densiﬂ ora

[

42 ] and P. patula [ 30 ] to increase the osmolarity of culture

media and ensure the success of the process ( see Note 4 ).

Germination stage (i.e., the conversion of somatic embryo s to

emblings) can be carried out at a broader range of concentra-

tions of Gelrite

® or other gellan gum brands.

4. Once medium is sterilized, it is supplemented with a source of

organic nitrogen that varies among Pinus species, being casein

hydrolysate plus l -glutamine in P. nigra [

47 ], P. strobus [ 39 ],

and P. sylvestris [

48 ], or EDM amino acid mixture [ 31 ] in P.

armandii [

44 ], P. densiﬂ ora [ 42 ], P. radiata [ 21 ], the most

commonly used ( see Note 5 ).

5. Plant growth regulator s ( PGR ) added to medium are as fol-

lows: (i) at initiation and proliferation stages, a cytokinin (ben-

zyladenine, BA) and an auxin (2,4-dichlorophenoxyacetic,

2,4-D) both at the concentration of 2.2 μM for P. strobus [

39 ],

2.7 and 4.5 μM, respectively, for P. radiata [

21 ], 4.4 and 13.6

μM for P. pinaster [

49 ] or the hybrid P. rigida x P. taeda [ 50 ],

2.2 and 9 μM for P. nigra [

27 ] ( see Note 6 ). Other hormones

Somatic Embryogenesis in Pine

408

can also be used for initiation and proliferation of EM , i.e.,

kinetin instead of BA (at 2.7 μM in P. halepensis [

22 ]) or in

combination with BA (both at 2.0 μM in P. taeda [

34 ]), and

auxins such as 1-naphthaleneacetic acid (NAA) instead of

2,4-D (at 10.7 μM in P. taeda [

34 ]) or in combination with

2,4-D (both at 4.5 μM in P. halepensis [

22 ]); (ii) at maturation

stage I (initial procedure of tissue resuspension), liquid medium

lacks PGR and organic nitrogen [

23 ]. For improving P. pinas-

ter [

37 ] and P. densiﬂ ora [ 42 ] maturation process, activated

charcoal (5 to 10 g/L) can be added to the liquid medium

used to resuspend EM ( see Note 7 ); at maturation stage II (tis-

sue culture on the ﬁ lter paper), abscisic acid ( ABA ) at a con-

centration ranging from 40 μM (in P. oocarpa [

46 ]) to 120 μM

(in P. strobus [

39 ]) is used.

6. The basal medium for germination phase is the same used for

the previous stages of the process, except in some species such

as P. radiata in which somatic embryo s are germinated in LP

medium [

51 ]. Germination medium lacks PGR and is usually

supplemented with sucrose . Sucrose concentration varies

depending on the species, e.g., at 10 g/L in P. densiﬂ ora [

42 ],

15 g/L in P. taeda [

52 ] and 30 g/L in P. radiata [ 23 ] and P.

halepensis [

22 ]. P. nigra germination culture medium contains

maltose (20 g/L), instead of sucrose [

47 ]. In species such as P.

armandii [

44 ], P. halepensis [ 22 ], P. nigra [ 47 ], or P. radiata

[

23 ], culture medium is supplemented with activated charcoal

to germinate the somatic embryos ( see Note 8 ).

7. The pH of culture media for all Pinus spp. is adjusted to

5.7–5.8.

8. For initiation, proliferation and germination , explants can be

cultured into 90 × 15 mm Petri dishes (20–25 mL of semisolid

medium), while for maturation the use of 90 × 20 mm Petri

dishes is recommended (40 mL of semisolid medium) ( see

Note 9 ).

3 Methods

1. Spray intact cones with 70 % (v/v) ethanol, split into quarters

and remove immature seeds. Use 10 % (v/v) H

2 O

2 plus two

drops of Tween 20

® for 8 min for sterilizing immature seeds,

then rinse three times with sterile distilled H

2 O under the ster-

ile conditions of a laminar airﬂ ow cabinet.

2. Excise out aseptically ( see Note 10 ) whole megagametophytes

containing immature embryos and place them horizontally

onto initiation medium ( see Note 11 ). Then, lay out cultures

in the growth chamber (Fig.

1a ) ( see Note 12 ).

3.1 Initiation

and Proliferation

Itziar Aurora Montalbán et al.

409

Fig. 1 ( a ) Initiation of EM in P. radiata megagametophytes cultured on EDM medium , bar 10 mm. ( b ) Proliferation

of EM of P. halepensis cultured on DCR medium , bar 6 mm. ( c ) P. radiata somatic embryo s obtained from

100 mg of EM cultured on EDM supplemented with 60 g/L sucrose and 60 μM ABA , bar 6 mm. ( d ) Tissue

overgrowth obtained from 150 mg of EM cultured on EDM supplemented with 60 g/L sucrose and 60 μM ABA,

bar 10 mm. ( e ) P. radiata somatic plantlets after 14 weeks germinating on half strength LP supplemented with

2 g/L activated charcoal , bar 12 mm. ( f ) Somatic P. halepensis plant growing in the greenhouse, bar 25 mm.

Somatic Embryogenesis in Pine

410

3. In most of the Pinus species tested, after 4–10 weeks on initia-

tion medium, proliferating EM with a size around 3–5 mm in

diameter is separated from megagametophytes. EM is subcul-

tured to proliferation medium every 2 weeks (Fig.

1b ). In

P. sylvestris [

28 ] and P. pinaster [ 37 ], in order to attain a high

amount of EM in a short period of time, weight 300 mg of EM

and resuspend it in liquid medium. Then pour it onto ﬁ lter-

paper disc and drain it using a Büchner funnel. Thereafter,

transfer the ﬁ lter paper with attached EM to proliferation

medium and subculture it to fresh medium each 2 weeks

( see Note 13 ).

4. During initiation and proliferation, keep cultures in darkness

[

21 ] or under low light intensity (5 μmol/m

2 /s [ 33 ]) at

21–24 °C [

41 ].

1. Maturation process is divided in two stages. For maturation

stage I, resuspend EM in liquid medium in 50 mL centrifuge

tubes ( see Note 14 ). Then, shake EM suspension vigorously by

hand for a few seconds.

2. 4–5 mL aliquot of the suspension, containing 300–500 mg of

EM , is used in P. brutia [

16 ] and P. strobus [ 53 ] and is poured

onto a ﬁ lter paper disc (Whatman no. 2, 70 mm) in a Büchner

funnel. For P. sylvestris [

48 ], EM amount can be decreased to

200 mg and, in species such as P. halepensis [

22 ], P. pinaster

[

37 ], or P. radiata [ 23 ], a low amount of EM (60–100 mg

fresh weight) is used to obtain the best results (Fig.

1c ) and

avoid overgrowth (Fig.

1d ) ( see Note 15 ).

3. Apply a vacuum pulse for 10 s. For maturation stage II, transfer

the ﬁ lter paper disc with attached EM to maturation medium

( see Note 16 ), such as in P. monticola [

38 ], P. nigra [ 47 ], or

P. taeda [

54 ]. On the contrary, in P radiata [ 23 ], P. pinaster

[

37 ], and P. sylvestris [ 48 ], the ﬁ lter paper discs with attached

EM are not subcultured through all maturation process.

4. During maturation, cultures are kept in darkness or under low

light intensity (5 μmol/m

2 /s), at 16-h photoperiod and

21–24 °C.

1. After 6–15 weeks, collect mature somatic embryo s, i.e., white

to yellowish, non-germinating somatic embryos with a distinct

hypocotyl region and at least three cotyledons (Fig.

1c ). A par-

tial desiccation pre- germination treatment has been described

in P. thunberghii , P. densiﬂ ora , P. armandii [

55 ], P. patula

[

30 ], P. nigra [ 47 ], and P. oocarpa [ 46 ]. Partial desiccation can

be carried out at 25 °C in a laminar airﬂ ow cabinet for 0–4 h

(fast method) or at high relative humidity, placing embryos

over 30 mm diameter ﬁ lter paper disks into a multiplate in

3.2 Maturation

3.3 Germination and

Acclimatization

Itziar Aurora Montalbán et al.

411

which some wells are ﬁ lled with 5–6 mL of sterile water, sealed

tightly and placed in darkness at 25 °C for 0–3 weeks (slow

method) [

55 ]. P. elliotii [ 56 ] somatic embryos can also be

stratiﬁ ed in order to increase somatic embryo conversion to

emblings ( see Note 17 ).

2. Culture somatic embryo s on Petri dishes with embryonal root

caps pointing downwards and tilt the Petri dishes vertically at

an angle of approximately 45–60°. In P. radiata [

23 ], P.

halepensis [

22 ] and P. nigra [ 47 ], the cultures are maintained

at 21–24 °C under a 16-h photoperiod at 40 μmol/m

2 /s for

1–2 weeks, and then at 120 μmol/m

2 /s provided by cool

white ﬂ uorescent tubes. In P. densiﬂ ora [

55 ], P. taeda , P. elliot-

tii , and P. palustris [

57 ], red wavelengths provided by light-

emitting diode (LED) improve somatic embryo germination .

3. After 6–8 weeks on germination medium, subculture the

plantlets once onto fresh germination medium. After another

6–8 weeks on germination medium, transfer the somatic plants

to trays with a sterile potting mix (Fig.

1e ). As potting mix, use

a peat:perlite, ratio 3:1 and 7:3 in P. radiata [

23 ] and P.

halepensis [

22 ], respectively, pine bark in P. patula [ 30 ], ver-

miculite in P. kesiya [

19 ], peat:vermiculite (3:1) in P. pinaster

[

37 ], and perlite:peat:vermiculite (1:1:1) in P. taeda [ 52 ] and

P. rigida x taeda [

50 ] ( see Note 18 ).

4. Acclimatize the plantlets in a greenhouse under controlled

conditions, decreasing humidity progressively [

8 ] ( see Note

19 ) (Fig.

1f ).

4 Notes

1. Ten megagametophytes per seed family or control cross should

be destructively sampled; the megagametophyte is carefully cut

longitudinally under an inverted microscope. Sometimes the

use of acetocarmine can help to see the zygotic embryo , espe-

cially at early stages of development [

21 ]. If the stage of most

zygotic embryos is not between early cleavage polyembryony

and ﬁ rst “bullet” stages with a dominant embryo [

33 ], initia-

tion rates of SE will be very low or zero.

2. To minimize high humidity and contamination, cones are

wrapped in ﬁ lter paper and stored in expanded polystyrene boxes.

3. For the development of somatic embryo s, it is necessary to

restrict water availability by physical or chemical means, such as

increasing osmotic agents (e.g., gellan gum or sugars)

concentration.

4. In this sense, some authors also add polyethylene glycol to

maturation medium [

16 , 30 , 42 , 44 , 46 ].

Somatic Embryogenesis in Pine

412

5. It is important to adjust the pH of thermolabile organic nitro-

gen solution (i.e., l -glutamine ) to 5.7–5.8.

6. The most commonly used PGR sources and concentrations are

those presented in Subheading

2.2 , but some authors use dif-

ferent plant growth regulator s for initiation such as N -(2-

chloro- 4-pyridyl)- N ′-phenylurea ( CPPU ) in P. pinaster [

58 ],

or PGR concentration is reduced to improve proliferation (in

P. densiﬂ ora [

42 ]).

7. When using a high concentration of activated charcoal in liq-

uid medium, it is important to adjust carefully pH of the

medium to 5.7–5.8. It is also important to shake liquid medium

before resuspending the EM , in order to avoid sedimentation

of activated charcoal.

8. Activated charcoal can be a critic factor for conversion of

somatic embryo s into plantlets, as proved in P. radiata [

23 ].

Thus, when trying to achieve conversion into plantlets of

somatic embryos in a speciﬁ c Pinus species for the ﬁ rst time, it

is suggested to test germination media with and without acti-

vated charcoal .

9. Semisolid media are prepared at least one week before being

used, while liquid medium is prepared the day before and

totally used within a week.

10. For this purpose we use Gerald forceps and scalpels (scalpel

blades number 11 or 20, depending on the size of the seeds).

11. Although some authors [

33 ] have increased initiation rates by

excising out immature zygotic embryo s from megagameto-

phytes (using a dissecting microscope), this procedure is time

consuming and requires sophisticated technical skills to avoid

damages or contamination of zygotic embryos.

12. It is advisable not to put more than ten explants per Petri dish,

in order to avoid later overlapping of extruding EM . Petri

dishes in all stages of the process are sealed with cling ﬁ lm.

13. Only peripheral parts of EM must be taken for proliferation,

and particularly for maturation of EM (it is recommended the

use of forceps). It is also important to be careful with the

amount of EM per aliquot; otherwise overgrowth of tissue on

the ﬁ lter paper would hinder development of somatic embryos

(Fig.

1d ). For this purpose, if SE has not been previously stud-

ied in a given species, it is convenient to test different amounts

of EM per aliquot.

14. Maturation is carried out once a sufﬁ cient amount of tissue is

achieved, usually after 4–8 subculture periods.

15. Sterilize ﬁ lter papers prior to use them for maturation process.

In order to avoid cross contaminations, use a different Büchner

funnel for each embryogenic cell line maturated; the use of

autoclavable plastic funnels makes this procedure easier.

Itziar Aurora Montalbán et al.

413

16. In some Pinus spp., such as P. sylvestris [ 48 ] and P. pinaster

[

37 ], the ﬁ lter paper with attached EM is subcultured fort-

nightly to fresh proliferation medium.

17. In P. radiata [

23 ], P. monticola [ 38 ], or P. halepensis [ 22 ] it is

not necessary to perform any pre- germination treatment if

somatic embryo s show a normal morphology.

18. These mixes can be supplemented with slow release osmocote

(at 750 g m

3 [ 38 ]). As suggested by several authors [ 41 ], fer-

tilization and pesticide treatments are the same as used for

seedlings, except that somatic seedlings are fertilized immedi-

ately after transplanting [

41 ].

19. The ﬁ rst 2 weeks after transplanting the plantlets, an acclimati-

zation tunnel is recommendable to maintain the humidity at

90–95 %.

Acknowledgments

This work was supported by Ministerio de Ciencia y Tecnología-

Spain (AGL2005-08214- CO2 -02) and Departamento de

Agricultura y Pesca-Basque Government (VEC2004021 and

VED2007014).

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Somatic Embryogenesis in Pine

417

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 22

Somatic Embryogenesis of Abies cephalonica Loud.

Jana Krajňáková and Hely Häggman

Abstract

Greek ﬁ r ( Abies cephalonica Loudon) belongs to the Mediterranean ﬁ r species and is widely distributed in

the mountains of Central and Southern Greece. Considering a climatic scenario, infestation by pathogens

or insects and ﬁ re episodes, it has been proposed that Mediterranean ﬁ rs could be in danger in some parts

of their present range but, on the other hand, could also replace other species in more northern zones with

temperate humid climates (e.g., silver ﬁ r, Abies alba Mill.). As ﬁ r species are generally highly productive

and therefore important for commercial forestry, they have traditionally been involved in conventional tree

improvement programs. A lot of effort has been put into the development of vegetative propagation meth-

ods for ﬁ rs, in order to rapidly gain the beneﬁ ts of traditional breeding to be utilized in reforestation. The

present paper provides up to date information on protocols for somatic embryogenesis (i.e., the most

promising in vitro method for vegetative propagation) of Greek ﬁ r. Moreover, the protocols for cryo-

preservation and long-term storage of embryogenic material are described as well.

Key words Cryopreservation , Ectomycorrhizal fungi , Fulvic acid s , Greek ﬁ r , Initiation , Maturation ,

Proliferation

1 Introduction

Greek ﬁ r ( Abies cephalonica Loudon) is a medium-sized tree,

widely distributed in mountains of Central and Southern Greece,

mainly at altitudes of 800–1700 m, covering an area of 200,000 ha

of productive and conservation forests [

1 ]. Most stands are rather

degraded and the present distribution is just a fraction of its poten-

tial natural area [

2 ]. The decrease has been attributed to various

reasons such as air pollution [

3 ], drought-related extreme periods,

infestation by mistletoe, pathogens or insects, root damage [

1 , 4 ,

5 ], ﬁ re episodes spreading over high altitudes [ 6 ], as well as the

effects of climate change [

7 ]. Considering a climatic scenario, it

has been proposed that Mediterranean ﬁ rs could be in danger in

some parts of their present range, but, on the other hand, could

also replace other species in more northern zones with temperate

humid climates (e.g. silver ﬁ r, Abies alba Mill) [

8 ]. As ﬁ r species are

generally highly productive and therefore important for

418

commercial forestry, they have traditionally been involved in con-

ventional tree improvement programs. A lot of effort has been put

into the development of vegetative propagation methods for ﬁ rs,

in order to rapidly gain the beneﬁ ts of traditional breeding to be

utilized in reforestation [

9 ]. Somatic embryo genesis (SE), i.e., the

development of embryos from somatic cells, with its potential for

mass multiplication has become a useful technique for large-scale

propagation of many coniferous species [

10 ]. In combination with

cryopreservation somatic embryo genesis makes it possible to pre-

serve important genotypes during ﬁ eld tests (reviewed in [

10 ]).

In Greek ﬁ r , like in other conifers, the multi-step regeneration

process of SE starts with induction of pro-embryogenic masses ,

followed by somatic embryo formation, maturation, desiccation

and plant regeneration (Fig.

1 ). Abies species were among the ﬁ rst

coniferous species where the induction of SE was reported [

11 ,

12 ]. However, a standard protocol for propagation by SE on a

large scale is still lacking, the only exception is SE of A. nordma-

nniana in which case the technology has already been tested in

large scale (Jens Find, personal communication). A. cephalonica

was regenerated by Krajňáková et al. [

13 ] and hybrid A. alba × A.

cephalonica was regenerated by Salajová et al. [

14 ]. Embryogenic

cultures of A. cephalonica and hybrid A. alba × A. cephalonica [

13 ,

14 ] have been derived from immature zygotic embryo s. In case of

hybrid ( A. alba × A. cephalonica ), initiation of embryogenic cul-

tures was achieved when using also mature embryos [

15 ] and coty-

ledons derived either from seedlings or somatic embryos (secondar y

or repetitive SE) [

16 ].

Somatic embryo genesis of several Abies species, including A.

cephalonica , differs from most of the other genera of the Pinaceae ,

because only cytokinin is needed for induction and proliferation

[

13 , 17 ]. Maturation of Greek ﬁ r and hybrid A. alba × A. cepha-

lonica somatic embryo s is promoted by abscisic acid and maltose is

the preferable carbohydrate. The addition of polyethylene glycol

promoted the development of somatic embryos [

15 , 18 ]. For ger-

mination , well-developed cotyledonary somatic embryos are

selected and subjected to a partial desiccation treatment for 3

weeks [

13 , 18 ]. Despite positive achievements, the bottlenecks in

A. cephalonica , like in most conifers, are the low initiation rate,

uneven maturation of embryos, problems in rooting and germina-

tion phases. This is due to poor understanding of embryo develop-

ment and therefore inability to develop proper SE methods for

Fig. 1 (continued) embryogenic cell mass and detail of proembryogenic cell masses after staining with aceto-

carmine and Evan’s blue ( f ). ( g ) Option for cryopreservation of the germplasm. ( h ) Maturation of somatic

embryos with embryogenic cell masses spread on ﬁ lter paper or ( i ) as clumps over solid medium (a cotyledon-

ary somatic embryo is showed in a small box ). ( j ) Plants prepared for experimental ﬁ eld trail

Jana Krajňáková and Hely Häggman

Fig. 1 Somatic embryo genesis of Abies cephalonica . ( a ) Elite tree of A. cephalonica . ( b ) Developing green cone,

shortly after meiosis. ( c ) Initiation of somatic embryo genesis using immature embryos and proliferation of

embryogenic cell mass , protruding from both sides of megagametophyte . ( d ) Initiation of somatic embryogen-

esis and proliferation of embryogenic cell masses ( arrow is pointing on the resin residuals). ( e ) Proliferating

420

practical purposes. Recently, the use of fulvic acid s, for improving

the proliferation abilities of Greek ﬁ r was studied [

19 ], as well as

the technique of cocultivation of ectomycorrhizal fungi with

embryogenic cell mass es which have led to improvements during

maturation [

20 ].

The aim of present paper is to provide up-to-date information

on protocols for Greek ﬁ r somatic embryo genesis, cryopreservation

of embryogenic cell mass es , and their long-term storage.

2 Materials

General equipment for tissue culture:

1. Laminar ﬂ ow hood.

2. Scalpels.

3. Forceps.

4. Growth chamber or cultivation room.

5. Autoclave.

6. pH-meter.

1. Seed cones containing immature seeds.

2. 70 % ethanol.

3. 4 % (w/v) CaOCl (Ca-hypochlorite).

4. Sterile distilled water.

5. 9 cm sterile Petri dishes.

6. 100 mL sterile beaker.

7. Initiation medium (Tables

1 and 2 ): Initiate either on solid

MS- based medium [

21 ] or SH [ 22 ] medium. Media are modi-

ﬁ ed as follows: Half-strength macroelement MS medium sup-

plemented with 20 g/L (58 mmol/L) sucrose , 1 mg/L (4.44

μmol/L) benzyl adenine (BA), 500 mg/L (3.4 mM) L -

GLUTAMINE , and solidiﬁ ed with 0.3 % (w/v) gellan gum

Phytagel ™ (Sigma) [

13 ]; SH medium , containing 20 g/L (58

mmol/L) sucrose, 1 mg/L (4.44 μmol/L) BA, 500 mg/L

(3.4 mM)

L - glutamine , and solidiﬁ ed with 0.3 % (w/v) gellan

gum Phytagel™ (Sigma) [

13 ].

8. Proliferation medium: MS-based initiation medium with addi-

tion of 0.1 % (w/v) casein hydrolysate [

13 ].

1. Sterile ﬁ lter paper discs.

2. Falcon tubes.

3. 9 cm Petri dishes.

2.1 In Vitro Protocols

for Somatic

Embryogenesis

2.1.1 Initiation, Induction,

and Proliferation

2.1.2 Maturation

Jana Krajňáková and Hely Häggman

421

Table 1

Concentrations of basic ingredients in half-strength macroelement MS medium [

21 ], SH medium

[ 22 ], and DCR medium [ 23 ] used for somatic embryo genesis of Greek ﬁ r

Component Half-strength macroelement MS SH DCR

Ingredient [mg/L] mM [mg/L] mM [mg/L] mM

Inorganic macro

NH

4 NO

3 825 10.3 – – 400 5

KNO

3 800 9.4 2500 25 334 3.3

Ca(NO

3 )

2 ·4H

2 O – – – – 543 2.3

CaCl

2 ·2H

2 O 220 1.5 200 1.4 84 0.57

MgSO

4 ·7H

2 O 185 0.75 400 1.6 370 1.5

KH

2 PO

4 85 0.625 – – 163 1.2

NH

4 H

2 PO

4 – – 300 2.6 – –

Inorganic micro

KI 0.83 0.005 1.0 0.006 0.8 0.005

H

3 BO

3 6.2 0.1 5.0 0.08 6.2 0.1

MnSO

4 ·4H

2 O 22.3 0.1 – –

MnSO

4 ·H

2 O – – 10 0.06 22 0.13

ZnSO

4 ·7H

2 O 8.6 0.030 1.0 0.0035 8.6 0.03

Na

2 MoO

4 ·2H

2 O 0.25 0.001 0.1 0.0004 0.24 0.001

CuSO

4 ·5H

2 O 0.025 0.0001 0.2 0.0008 0.25 0.001

CoCl

2 ·6H

2 O 0.025 0.0001 0.1 0.0004 0.024 0.0001

NiCl

2 ·6H2O – – – – 0.024 0.0001

Na

2 -EDTA 37.25 0.1 20 0.055 34 0.1

FeSO

4 ·7H

2 O 27.85 0.1 15 0.055 27.8 0.1

Organics

Myoinositol 100 0.555 1000 5.55 198 1.1

Nicotinic acid 1.0 0.0812 5.0 0.41 0.5 0.0041

Pyridoxine-HCl 1.0 0.0048 0.5 0.0024 0.5 0.0024

Thiamine- HCl 1.0 0.003 5.0 0.015 1 0.003

Glycine 2.0 0.0266 – – 2.0 0.0266

L-glutamine 500 3.42 500 3.42 248 1.7

Somatic Embryogenesis of Abies cephalonica

422

Table 2

Composition of the initiation, proliferation, maturation, and conversion media during cultivation of Greek ﬁ r embryogenic cultures [ 13 , 18 , 24 ]

Medium composition Initiation [ 13 ]

Proliferation

[ 13 , 24 ]

Liquid medium for

suspension [ 24 ] Maturation Conversion

Inorganics and organics

Half- strength

macroelement

MS SH

Half-strength

macroelement

MS

Half-strength

macroelement

MS

DCR

[ 13 ]

Half-strength

macroelement

MS [ 24 ]

Half-strength

macroelement

MS [ 24 ]

Half- strength

DCR [ 13 , 18 ]

Casein hydrolysate (g/L) 1 1 0.5 0.5 0.5

L - Glutamine 0.5 0.5 0.25 0.25 0.25

Sucrose (g/L) 20 20 20 20

Maltose (g/L) 30 30 30 20

BA (mg/L) 1 1

ABA (mg/L) 8.5 17 8.5

Phytagel (g/L) 3 3 3 2.5 2.5 2.5

Agar (g/L) 10

PEG 4000 (g/L) 10

Jana Krajňáková and Hely Häggman

423

4. Liquid proliferation media without plant growth regulator s

(Table

2 ) for making suspension with embryogenic cell mass es

( ECMs ) [

24 ].

5. Perform maturation either on solid DCR medium : DCR with

8.5 mg/L (32 μM) abscisic acid ( ABA ), 10 % (w/v) polyethyl-

ene glycol ( PEG ) 4000, 0.05 % (w/v) casein hydrolysate , 250

mg/L, 1.7 nmol/L

L -glutamine , 30 g/L (83.3 mM) maltose

[

13 ] ( see Note 1 ) or solid MS media: (a) half-strength

macronutrient MS medium with 17 mg/L (64 μM) ABA, 10

% (w/v) polyethylene glycol (PEG) 4000, 0.05 % (w/v) casein

hydrolysate, 250 mg/L (1.7 nmol/L)

L - glutamine , 30 g/L

(83.3 mM maltose) [

24 ]; (b) half-strength macronutrient MS

medium with 8.5 mg/L (32 μM) ABA, 0.05 % (w/v) casein

hydrolysate, 250 mg/L (1.7 nmol/L)

L -glutamine, 30 g/L

(87.6 mM) maltose [

24 ] (Tables 1 and 2 ).

1. 150 mL tissue culture jars.

2. Solid DCR medium for conversion : Half-strength DCR

hormone- free medium with 20 g/L (58 nmol/L) maltose

[

13 , 18 ] (Tables 1 and 2 ).

3. Non-fertilized horticultural peat and perlite.

4. Plastic containers, commercial fertilized peat (VAPO, Finland)

with 1 kg/L basic fertilizer: 9.7 % N, 7.5 % P, 14.4 % K, 5.0 %

Ca, 6.6 % S, 3.8 % Mg, 0.27 % Fe, 0.13 % Mn, 0.04 % B, 0.05

% Zn, 0.25 % Cu, and 0.09 % Mo and 3 kg/L limestone dust

with Mg, commercial 0.2 % 5-Superex fertilizer (Kekkilä,

Finland).

2.1.3 Conversion

and Acclimatization to Ex

Vitro

Table 3

Composition of proliferation and pretreatment media for cryopreservation of Greek ﬁ r embryogenic

cell mass es [ 13 , 24 , 25 ]

Medium

composition Proliferation Pretreatment Cryo-treatment

Inorganics and

organics

Half-strength

macroelement MS

Half-strength

macroelement MS

Half-strength

macroelement MS

Half-strength

macroelement MS

Casein hydrolysate

(g/L) 1 0.5 0.5 1

L - Glutamine 0.5 0.25 0.25 0.5

Sucrose (g/L) 20 68.5 137 137

BA (mg/L) 1 1 1

Phytagel (g/L) 3 3 3

Somatic Embryogenesis of Abies cephalonica

424

1. Cryovials and markers.

2. Cryobox or cryocanes for immersion of the cryovials in liquid

nitrogen ( LN ).

3. Sterile tips for pipets of different volumes (0.2 μL to 1 mL).

4. Programmable controlled-temperature chamber or Nalgene™

freezing container and isopropanol.

5. Ice.

6. Dewar for the conservation of samples in LN .

7. Actively proliferating embryogenic cell mass es (10- to 12-day

old, after the last regular transfer).

8. Solid MS based medium for cryopreservation: (a) half-strength

macroelement MS medium , hormone-free, containing 68.5

g/L (0.2 M) sucrose ; (b) half-strength macroelement MS

medium, hormone-free, containing 137 g/L (0.4 M sucrose)

[

24 , 25 ] (Tables 1 and 3 ).

9. Liquid MS-based medium for cryopreservation: Half-strength

macroelement MS medium , hormone-free, containing 137

g/L (0.4 M) sucrose [

24 , 25 ] (Tables 1 and 3 ).

10. Solid MS-based proliferation medium: Half-strength macroel-

ement MS medium with 20 g/L (58.4 mM) sucrose , 1 mg/L

(4.44 μmol/L) BA, 500 mg/L (3.4 mM)

L -glutamine , 0.1 %

(W/v) casein hydrolysate [

13 , 24 , 25 ] (Tables 1 and 3 ).

11. PGD cryoprotectant solution: 10 % PEG 6000, 10 % glucose,

10 % dimethyl sulfoxide ( DMSO ) in H

2 O, ﬁ lter sterilized.

3 Methods

1. Solid MS, SH, and DCR culture media (Tables 1 and 2 ) for

initiation, proliferation, and maturation are prepared in 9 cm

Petri dishes and liquid media to arrest proliferation in 250 mL

Erlenmeyer ﬂ asks. The pH of medium is adjusted to 5.7 prior

adding the solidifying agent. Media for conversion are pre-

pared in tissue culture jars (Magenta vessels). Aqueous stock

solutions of

L -glutamine and ABA are ﬁ lter sterilized and added

to the medium after autoclaving. Separately autoclaved poly-

ethylene glycol is mixed with the rest of the maturation medium

in laminar ﬂ ow hood to get the ﬁ nal volume.

2. Immature zygotic embryo s surrounded by the megagameto-

phyte (called immature zygotic embryos) and isolated from

immature seed cone are most favorable material for initiating

SE of Greek ﬁ r . The optimum developmental stage of imma-

ture zygotic embryos for initiation is the precotyledonary stage

(i.e., 1 month after fertilization but before the formation of

cotyledons) [

13 ] ( see Note 2 ) (Fig. 1a, b ). However, in case of

2.2 Cryopreservation

3.1 Somatic

Embryogenesis

3.1.1 Culture Media

Preparation, Explant

Excision and Sterilization,

and Culture Initiation

Jana Krajňáková and Hely Häggman

425

hybrid A. alba × A. cephalonica , also mature zygotic embryos

are used [

15 ] as well as cotyledons from seedlings and emblings

[

16 ] ( see Notes 3 and 4 ).

3. Immature seed cones are rinsed with 70 % ethanol for 2 min,

after which immature seeds are removed from the cones using

scalpels and forceps and placed in sterile beaker with sterile

distilled water.

4. Seeds are surfaced sterilized for 20 min in 4 % (w/v) CaOCl,

and rinsed three times for 5 min with sterile distilled water ( see

Note 5 ).

5. Seed coats are opened and removed with forceps and imma-

ture zygotic embryo s surrounded by megagametophytes are

excised and placed onto MS or SH medium for initiation ( see

Note 6 ) (Fig.

1c ).

6. Immature zygotic embryo s are ﬁ rst cultured for 4 weeks, and

thereafter transferred onto new media for an additional 4

weeks. However, it is recommended to control the contamination

problems within the ﬁ rst week of cultivation. The contaminated

immature embryos should be discarded and not contaminated

transferred to a fresh medium ( see Note 7 ) (Fig.

1d ).

7. Initiation and induction is performed in the dark at 22 ± 2 °C.

1. Embryogenic cell masses start to protrude from different parts

(micropylar end being the most frequent) of the responsive

explants (immature zygotic embryo s, surrounded by megaga-

metophytes) 4–6 weeks after initiation (Fig.

1c ). Embryogenic

tissues are excised from each explant separately (each one rep-

resenting one genotype) and transferred to a new Petri dish

with proliferation medium (MS or SH) to form a new cell line.

2. To maintain the proliferation of ECMs , they are transferred to

fresh medium every 3 weeks. ECMs can be subcultured for sev-

eral months ( see Note 8 ) (Fig.

1e–g ) in the dark at 22 ± 2 °C.

3. ECMs can be used as such for maturation.

1. Clumps of ECMs (or ﬁ lter paper covered by a thin layer of

ECM suspension) are transferred to maturation medium 1

(Table

2 ; see Notes 9 and 10 ) for the ﬁ rst 6 weeks, followed by

regular transfers to fresh media at 2-week intervals (Fig.

1h , i ).

For further development of somatic embryo s, ABA concentra-

tion is decreased and PEG -4000 is omitted from the medium

(Table

2 ).

2. For preparing the suspension, 4 g of fresh ECM is transferred

to sterile Falcon ﬂ asks with 20 mL of liquid hormone-free pro-

liferation medium (Table

2 ). Suspension is gently mixed by

3.1.2 Proliferation

of Embryogenic Cultures

3.1.3 Maturation

of Embryogenic Cultures

Somatic Embryogenesis of Abies cephalonica

426

vortex and allowed to settle. After the removal of supernatant,

1 mL of suspension, containing approximately 250 mg ECM

(fresh weight), is plated onto sterile Whatman ﬁ lter paper

placed on maturation medium [

24 ].

3. Maturation is performed in the dark at 22 ± 2 °C.

1. Mature healthy cotyledonary somatic embryo s are carefully

detached from the embryogenic cell mass es (Fig.

1I , small

box) and transferred on empty Petri plates (diameter ca. 4 cm)

which are placed into bigger Petri plates (diameter ca. 9 cm)

with sterile distilled water for 3 weeks desiccation period.

2. During the desiccation period, somatic embryo s are stored in

the dark, at the temperature of 4 °C.

3. Afterwards, desiccated embryos are placed onto the hormone-

free half-strength DCR medium with 20 g/L (58 mM) malt-

ose , solidiﬁ ed with 1 % (w/v) agar [

13 , 18 ] (Tables 1 and 2 ).

4. The base of somatic embryo s is gently inserted into the

medium.

5. Somatic embryo s are germinated at the temperature of 22 ± 2

°C and the light intensity is kept for the ﬁ rst 2 weeks at 30 μE/

m

2 /s (16 h photoperiod), and then gradually augmented up to

75 μE/m

2 /s.

6. Somatic embryo -derived plantlets are carefully detached from

the medium and roots are washed. Thereafter plantlets are

planted into small plastic greenhouses containing non-fertil-

ized horticultural peat and perlite ( v:v ) (2:1). For the ﬁ rst 2

weeks the plantlets are kept under mist in order to keep relative

humidity at approximately 90 %, after which the humidity is

gradually decreased.

7. After 1 month the plantlets are transferred into bigger contain-

ers (diameter ca. 5–6 cm) containing commercial fertilized

peat (VAPO, Finland) in a greenhouse. During the growing

season, plantlets are fertilized monthly with commercial 0.2 %

5-Superex fertilizer (Kekkilä, Finland) (Fig.

1j ).

1. Transfer actively proliferating ECMs , size 300 ± 50 mg, on

MS- based proliferation medium and cultivate at 5 °C in the

dark for 14 days ( see Note 11 ).

2. After cold hardening, transfer the culture onto proliferation

medium supplemented with 0.2 M sucrose for 24 h, and after-

wards onto 0.4 M sucrose medium for another 24 h.

3. Transfer about 3–4 ECM clumps into 400 μL of hormone-free

proliferation medium ( item 9 , Subheading

2.2 ) which is added

into 2 mL cryotubes on ice.

4. Add PGD cryoprotective solution dropwise over a period of

30 min to give a ﬁ nal concentration of 5 %.

3.1.4 Desiccation-

Conversion of Somatic

Embryos

and Acclimatization

to Ex Vitro

3.2 Cryopreservation

by Controlled- Rate

Cooling

3.2.1 Cold Hardening,

Pretreatments,

and Cryostorage

Jana Krajňáková and Hely Häggman

427

5. Leave the cryotubes to stand for 2 h on ice before freezing.

6. After ﬁ nishing the cryoprotection phase, freeze the samples at

a rate of 10 °C/h (0.17 °C/min) to the prefreezing tempera-

ture of −38 °C, using a programmable controlled-rate freezer

( see Note 12 ).

7. After reaching the terminal temperature, immerse the cryo-

tubes containing samples in LN and store.

1. Thaw the cryovials in a 37 °C water bath and then transfer

them on ice.

2. Rinse the surfaces of cryovials with 70 % ethanol. Pay attention

to labeling.

3. Plate (dispense) the contents of the cryovials on an autoclaved

ﬁ lter paper disc, placed on proliferation medium in a 90 mm

Petri dish with 0.4 M sucrose . Incubate cultures for 1 h.

4. After 1 h, transfer ﬁ lter papers with suspensions onto fresh pro-

liferation medium with 0.2 M sucrose and incubate for 24 h in

the dark at 22 ± 2 °C.

5. After 24 h, transfer ﬁ lter papers with suspensions on the prolif-

eration medium (Tables

1 and 2 ).

6. Examine the viability of cells by staining the suspension culture

with 0.5 % FDA (ﬂ uorescein diacetate) and observe at the

microscope under UV light.

7. Monitor cultures regularly and transfer them onto fresh prolif-

eration medium at 2-week intervals ( see Note 13 ).

8. After observing the recovery (i.e., new proliferation growth),

transfer the embryogenic cell masses on fresh proliferation

medium without ﬁ lter paper disc.

9. Embryo maturation is established when proliferation of cryo-

preserved ECMs is comparable to non-cryopreserved cultures

( see Note 14 ).

4 Notes

1. DCR medium was originally used for tissue cultures of Douglas

ﬁ r [ Pseudotsuga menziesii (Mirb.) Franco] [

23 ] and have been

used for cultivation of other coniferous species, such as Pinus

[

26 ] and Abies species [ 13 , 14 ].

2. The cones with immature zygotic embryo s can be collected

and stored at 4 °C for at least 2 months without losing the abil-

ity to induce somatic embryo genesis [

13 , 27 ].

3. Mature embryos, used for initiation of SE, were excised from

hybrid seeds of A. alba × A. cephalonica , stored from 6 months

3.2.2 Thawing

and Recovery

Somatic Embryogenesis of Abies cephalonica

428

to 4 years [ 15 ]. Embryos isolated from seeds stored for 6

months showed 27 % initiation frequencies, and those isolated

from 1-year stored seeds 29 %. Embryos from seeds stored for

4 years did not response.

4. Embryogenic cultures have been initiated on cotyledon

explants dissected from seedlings or emblings of the hybrid A.

alba × A. cephalonica [

16 ]. Cotyledons of seedling origin gave

relatively low initiation frequencies (about 2 %). In embling-

derived cotyledons, the initiation was cell-line dependent and

reached values between 1 and 24 %.

5. Due to the toxic nature of HgCl

2 (0.1 %, w/v, solution used),

this sterilizing agent was omitted from tissue culture protocols.

However, it was successfully used when applied to immature

and mature seeds of A. cephalonica [

13 ] and A. alba × A. cepha-

lonica hybrid [

15 ]. Positive results were also obtained with

15 % H

2 O

2 [ 14 ].

6. There were no signiﬁ cant differences in initiation frequencies

when half-strength MS and SH media were compared [

13 ].

DCR and LM media have also been used for induction of

somatic embryo genesis from hybrid immature zygotic embryo s

of A. alba × A. cephalonica . However, these media turned out

to be impropriate for initiation of embryogenic cell masses

[

14 ]. On the other hand, ECMs were induced on cotyledon

explants isolated from emblings and seedlings of A. alba × A.

cephalonica hybrid on DCR-based medium [

16 ].

7. Seeds of A. cephalonica , like other Abies species, are full of res-

ins. Sometimes these resins are transferred during isolation of

embryos onto initiation medium and they create white plaques

similar to bacterial contamination. When observing this phe-

nomenon it is recommended to cultivate “suspicious” Petri

plates overnight at higher temperature (37 °C), as well as to

transfer the culture on a bacterial cultivation medium. If the

plaques remain of the same size, they are resins; if they grow,

then it is bacterial contamination.

8. During prolonged proliferation, the regeneration ability of

ECMs decreases. It is therefore very important to start with

cryopreservation when stable proliferation is achieved.

9. A suspension made with embryogenic cell mass es can also be

used for maturation [

19 , 24 , 27 , 28 ].

10. Arrest of proliferation can be achieved either on solid or liquid

hormone-free half-strength macroelement MS medium [

24 ,

27 , 29 ].

11. Step of cold-hardening has been omitted from cryo-

preservation protocol without noticing the decrease in viability

of cultures after thawing, however only two cell lines were

tested [

30 ].

Jana Krajňáková and Hely Häggman

429

12. Nalgene Freezing Container, Mr. Frosty, ﬁ lled with isopropa-

nol alcohol, was successfully used for cooling down the cryovi-

als, instead of using a programmable controlled-rate freezer

[

30 ].

13. The most precise way of monitoring the new proliferation is

determination of proliferation ratio ( w

0

/ w

i

) in which w

i

is the

initial fresh weight of sample after thawing and w

0

is the weight

at the time of subculturing, generally 2, 4, or 6 weeks after

thawing [

24 , 30 ].

14. Successful cryopreservation was published also for the ECMs

of ﬁ r hybrids ( A. alba × A. cephalonica , A. alba × A. numidica )

with pre-culturing on media with 0.4 or 0.8 M sorbitol for 24,

48, or 72 h and addition of 5 % (v/v) DMSO as a cryoprotec-

tant [

31 ].

Acknowledgments

The authors are grateful for the working facilities provided by

University of Oulu, Genetics and Physiology Department.

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 23

Somatic Embryogenesis in Horse Chestnut

( Aesculus hippocastanum L.)

Maurizio Capuana

Abstract

Embryogenic cultures of horse chestnut ( Aesculus hippocastanum L.) can be obtained from different

organs and tissues. We describe here the induction from stamen ﬁ laments and the procedures applied for

the successive phases of somatic embryo development and maturation. Embryogenic tissues are obtained

on Murashige and Skoog medium containing 9.0 μM 2,4-dichlorophenoxyacetic acid. Somatic embryos

develop after transfer to hormone-free medium enriched with glutamine. Maturation and germination of

isolated embryos are achieved by transfer to medium containing polyethylene glycol 4000 and activated

charcoal, successive desiccation treatment, and cold storage at 4 °C for 8 weeks.

Key words Anther ﬁ lament , Conversion , Desiccation , Maturation , Plant growth regulator s

1 Introduction

Common horse chestnut ( Aesculus hippocastanum L.) is one of the

12 species of the genus Aesculus , family Hippocastanaceae, that

comprises deciduous trees and shrubs distributed in the Northern

Hemisphere. There are two Eurasian species, A. hippocastanum

and A. chinensis , var. chinensis , both commonly used in medicine.

From A. hippocastanum , bark, leaves, and seed extract (HCSE)

have been used for several medical treatments [

1 ]; among its natu-

ral compounds, aescin, a saponin mixture extracted from this spe-

cies, displays diverse activities, including anti-inﬂ ammatory,

antiviral, and antioxidative properties [

2 ]. Since the beginning of

this century, the number of horse chestnuts with bleeding cankers

has increased in Europe [

3 ], highlighting the need to accelerate

the release of tolerant genotypes. Vegetative propagation of

selected superior trees is, thus, important for both environmental

and industrial purposes [

4 ]. Ornamental forms of horse chestnut

are generally multiplied by grafting or cuttings [

5 ]. In vitro propa-

gation methods have the advantage of speeding up the multiplication

432

process and embryogenesis, in particular, has a huge productive

potential to be exploited.

Somatic and gametic embryogenesis have been obtained from

different primary explants of horse chestnut , such as microspores

[

6 ], anther ﬁ lament s [ 7 ], zygotic embryo s [ 8 ], leaf segments [ 9 ],

and stem explants [

10 ]. Embryogenic tissues may also be used for

long-term conservation by cryopreservation [

11 ]. In this chapter,

a protocol for the induction and development of somatic embryo s

from ﬂ ower ﬁ laments is described. Compared to other kinds of

explants, this material offers the advantage of a lower presence

of contaminants and, consequently, an easier in vitro establish-

ment of culture.

2 Materials

1. Collect ﬂ ower buds before their opening and preferably from

the outer part of the crown of the selected plant(s), where the

lower humidity conditions promote the collection of healthier

explants; store them at 4 °C until use ( see Note 1 ).

1. Murashige and Skoog salts and vitamins (MS) [

12 ]; Woody

Plant Medium salts and vitamins (WPM) [

13 ] (Table 1 ).

2.

D -Sucrose pure.

3. Agar (B&V, Italy).

4. 2,4-Dichlorophenoxyacetic acid (2,4-D).

5. N

6 -benzyladenine (BA).

6. Indole-butyric acid (IBA).

7. 0.1 and 1.0 M KOH solutions.

8. 0.1 and 1.0 M HCl solutions.

9. Glutamine .

10. Polyethylene glycol 4000 ( PEG ).

11. Activated charcoal ( AC ).

12. 125, 500 mL glass ﬂ asks.

13. 500, 1000 mL beakers.

14. 500, 1000 mL cylinders.

15. Sterile Petri dishes (90 mm in diameter).

16. Tissue culture facilities: Precision balance, magnetic stirrer,

magnetic bars, microwave cooker, pH meter, autoclave for

sterilization, forceps, scalpels, sterilizer, laminar ﬂ ow bench,

growth chamber, refrigerator.

1. Tap water.

2. Ethanol (70 %).

2.1 Plant Material

2.2 Preparation

of Culture Media

2.3 Explant

Sterilization

Maurizio Capuana

433

3. Sodium hypochlorite solution (bleach solution at 7 g/L active

chlorine).

4. Distilled water (autoclaved reverse-osmosis water).

5. 125, 250 mL sterilized glass ﬂ asks.

6. 50, 100, 250 mL cylinders.

7. Tissue culture facilities: Forceps, sterilizer, laminar ﬂ ow bench,

growth chamber, refrigerator.

Table 1

Plant culture media: formulations of Murashige and Skoog (MS, [ 12 ]) and

Lloyd and McCown (WPM, [ 13 ])

MS (mg/L) WPM (mg/L)

KNO

3 1900 –

NH

4 NO

3 1650 400

MgSO

4 ·7H

2 O 370 370

KH

2 PO

4 170 170

CaCl

2 ·2H

2 O 440 96

Ca(NO

3 )

2 ·4H

2 O – 556

K

2 SO

4 – 990

H

2 BO

3 6.2 6.2

MnSO

4 ·4H

2 O 22.3 22.3

ZnSO

4 ·7H

2 O 8.6 8.6

Na

2 MoO

4 ·2H

2 O 0.25 0.25

CuSO

4 ·5H

2 O 0.025 0.25

CoCl

2 ·6H

2 O 0.025 –

KI 0.83 –

FeSO

4 ·7H

2 O 27.8 27.8

Na

2 EDTA·2H

2 O 37.3 37.3

Sucrose 20,000 20,000

Glycine 2.0 2.0

Pyridoxine·HCl 0.5 0.5

Nicotinic acid 0.5 0.5

Thiamine· HCl 0.1 1.0

Myo-inositol 100 100

Horse Chestnut Somatic Embryogenesis

434

1. Precision balance.

2. Magnetic stirrers, magnetic bars.

3. Microwave cooker.

4. pH meter.

5. Autoclave for sterilization.

6. Laminar ﬂ ow bench.

7. Refrigerator.

1. Plastic trays (with 3.5 cm diameter holes).

2. 6–8 cm diameter plastic pots.

3. Potting medium (garden soil, peat, sand, 3:1:1 by volume).

4. Greenhouse equipped with “mist” system.

3 Methods

It is possible to induce somatic embryo genesis on different kinds

of explants, such as mature or immature zygotic embryo s, portions

of leaves, and ﬂ ower parts. In this chapter we describe the induc-

tion of somatic embryogenesis from anther ﬁ lament s. Using these

explants we can start a clonal propagation cycle from a material of

identiﬁ able genetic origin, allowing the mass propagation of plants

selected for superior traits (shape, vigor, pest and insect resistance,

stress adaptability, etc.).

The following protocol, based on the experiences of different

authors [

14 – 17 ], comprises the following stages: (1) culture

media preparation; (2) plant material collection and sterilization;

(3) somatic embryo genesis induction; (4) somatic embryo devel-

opment ; (5) somatic embryo maturation ; (6) somatic embryo

conversion ; and (7) plantlet acclimatization.

1. Prepare MS and WPM media (Table

1 ) in double- distilled

water, supplemented with 2 % sucrose . Store at 4 °C.

2. Prepare 2,4-D, BA, and IBA stock solution: 2,4-D must be

dissolved in a few drops of absolute ethanol. For BA, use 1.0 M

KOH. Store at 4 °C.

3. Induction medium: Use MS supplemented with 9 μM 2,4-D.

4. Embryo development medium: Use plant growth regulator

( PGR )-free MS medium , containing 400 mg/L glutamine

(ﬁ lter-sterilized).

5. Maturation medium: Use PGR -free MS medium , containing

50 mg/L PEG and 1 g/L AC .

2.4 Laboratory

Equipment

2.5 Acclimatization

of Plantlets

3.1 Culture Media

and Conditions

Maurizio Capuana

435

6. For conversion , apply a slow- desiccation procedure by placing

the mature somatic embryo s, contained in empty and non- sealed

Petri dishes, on the laminar ﬂ ow bench and leave the material

under the air ﬂ ow for 48 h. Then, transfer somatic embryos to

conversion medium: WPM supplemented with 2 % sucrose ,

0.7 % agar , 0.2 mg/L BA and 0.02 mg/L IBA.

7. Adjust the pH of the media to 5.6 using HCl or KOH (1.0 and

0.1 M).

8. Add agar (0.7 %).

9. Sterilize the media by autoclaving at 121 °C and 108 kPa for

20 min.

10. Store the autoclaved media at 4 °C for a maximum of 60 days.

1. Cut the ﬁ laments and rinse them under slow running tap water

for 1 h.

2. Sterilize the laminar ﬂ ow surface by 70 % ethanol before use.

3. Disinfect the ﬁ laments by soaking in 70 % ethanol solution for

2 min, followed by two 2-min rinses in sterile distilled water;

disinfect again by soaking in 20 % sodium hypochlorite (1.4 %

active chlorine) solution for 20 min, with three ﬁ nal rinses in

sterile distilled water, under the laminar air ﬂ ow and using ster-

ilized glass ﬂ asks.

1. Under the laminar ﬂ ow bench, pick up the ﬁ laments and place

them horizontally on the induction medium (20–25 ﬁ laments

per Petri dish).

2. Incubate the cultures in the growth room (or cabinet) in

darkness.

3. After 1 month, transfer the explants onto fresh induction

medium.

1. After 1 month, transfer explants with emerging embryogenic

tissues from the induction medium to a PGR -free MS medium

containing 400 mg/L of ﬁ lter-sterilized glutamine ( embryo

development medium).

2. Incubate cultures at 16-h photoperiod light condition, 60.0

μmol/m

2 /s photosynthetically active radiation, for 1 month.

3. Subculture the material at 4-week interval ( see Note 2 ).

1. From clusters of maturing somatic embryo s (at stage from

globular to torpedo, before the developmental phase showed

in Fig.

1b ), isolate globular embryos (“singularization”) and

culture them for 4 weeks on MS medium containing 50 g/L

PEG and 1 g/L AC (maturation medium) ( see Note 3 ).

3.2 Explant Surface

Sterilization

3.3 Somatic

Embryogenesis

Induction

3.4 Somatic Embryo

Development and

Embryogenic Tissue

Proliferation

3.5 Somatic Embryo

Maturation

and Conversion

Horse Chestnut Somatic Embryogenesis

436

Fig. 1 ( a ) Embryogenic tissues protruding from the anther ﬁ lament of horse chestnut . ( b ) Cluster of maturing

somatic embryo s: most embryos are at cotyledonal stage, many others are at stages from globular to torpedo.

( c ) Converting somatic embryos. ( d ) Plantlet ready for transplant to pot and acclimatization

Maurizio Capuana

437

2. For conversion , apply a slow- desiccation procedure, by placing

the mature somatic embryo s, contained in empty and non-

sealed Petri dishes, on the laminar ﬂ ow bench and leave the

material under the airﬂ ow for 48 h (Fig.

1c ; see Note 4 ).

3. Transfer somatic embryo to PGR -free MS medium and store

cultures at 4 °C in darkness for 8 weeks.

4. Transfer somatic embryo s to conversion medium for a 4-week

period.

1. Select the converted somatic embryo s (i.e., with developing

apical pole and roots) and wash the roots under running tap

water to remove the adhering solidiﬁ ed culture medium.

2. Insert the plantlets in 35 mm diameter trays, ﬁ lled with potting

mixture and place the trays on a greenhouse bench equipped

with a mist system for 3–4 weeks.

3. Move the trays to a non-misted bench under a tunnel covered

with plastic foil, where they remain for about 3 weeks to allow

a gradual transition to ambient atmosphere.

4. Transplant the plantlets to larger pots (60–80 mm diameter)

and place the pots in a shaded area of the nursery fur further

growth.

4 Notes

1. Following the indication of Radojevic [ 5 ], ﬂ ower buds must be

collected at stage 3–4, when the buds (2–3 mm in length) are

completely closed.

2. Generally, embryogenic tissues continue to proliferate after

transfer to PGR -free medium This material can be sub-cul-

tured for years showing a very slow decline of proliferation

capacity; it is advisable, however, to transfer the cultures onto

a BA- containing medium (4.4 μM) after some months of sub-

culturing on PGR-free medium.

3. An asynchronous development of somatic embryo s may be

observed in every phase of maturation (Fig.

1b ). It is frequent

the development of irregular embryos showing hypertrophy,

absence of a well-organized shoot meristem, abnormal cotyle-

don shapes, or more than two cotyledons.

4. Somatic embryo conversion in horse chestnut can be

problematic. Better results can be achieved if, before applying

the desiccation procedure, somatic embryo s, as illustrated

above, are cultured for 4 weeks on medium containing PEG

(50 g/L) in combination with AC (1 g/L).

3.6 Acclimatization

Horse Chestnut Somatic Embryogenesis

438

References

1. Sirtori CR (2001) Aescin: pharmacology,

pharmacokinetics and therapeutic proﬁ le.

Pharmacol Res 44:183–193

2. Pittler MH, Ernst E (2012) Horse chestnut

seed extract for chronic venous insufﬁ ciency.

Cochrane Database Syst Rev 11:CD003230

3. Schmidt O, Dujesiefken D, Stobbe H, Moreth

U, Kehr R, Schröder T (2008) Pseudomonas

syringae pv. aesculi associated with horse chest-

nut bleeding canker in Germany. For Pathol

38:124–128

4. Oleksyn J, Kloeppel BD, Lukasiewicz S,

Karolewski P, Reich PB (2007) Ecophysiology

of horse chestnut ( Aesculus hippocastanum L.)

in degraded and restored urban sites. Pol J Ecol

55:245–260

5. Radojevic L (1991) Horse chestnut ( Aesculus

spp.). In: Bajaj YPS (ed) Biotechnology in agri-

culture and forestry, Tree III, vol 16. Springer,

Heidelberg, pp 111–141

6. Radojevic L (1978) In vitro induction of

androgenic plantlets in Aesculus hippocastanum

L. Protoplasma 96:369–374

7. Jörgensen J (1989) Somatic embryogenesis in

Aesculus hippocastanum L. by culture of ﬁ la-

ment callus. J Plant Physiol 135:240–241

8. Profumo P, Gastaldo P, Bevilacqua L, Carli S

(1991) Plant regeneration from cotyledonary

explants of Aesculus hippocastanum L. Plant Sci

76:139–142

9. Dameri RM, Caffaro I, Gastaldo P, Profumo P

(1986) Callus formation and embryogenesis

with leaf explants of Aesculus hippocastanum

L. J Plant Physiol 126:93–96

10. Gastaldo P, Carli S, Profumo P (1994) Somatic

embryogenesis from stem explants of Aesculus

hippocastanum . Plant Cell Tiss Org Cult 39:

97–99

11. Lambardi M, De Carlo A, Capuana M (2005)

Cryopreservation of embryogenic callus of

Aesculus hippocastanum L. by vitriﬁ cation/

one-step freezing. Cryo Letters 26(3):

185–192

12. Murashige T, Skoog FA (1962) A revised

medium for rapid growth and bioassays with

tobacco tissue cultures. Physiol Plant 15:

473–497

13. Lloyd G, McCown BH (1980) Commercially-

feasible micropropagation of mountain laurel,

Kalmia latifolia , by use of shoot tip culture.

Proc Int Plant Prop Soc 30:421–427

14. Radojevic L (1988) Plant regeneration of

Aesculus hippocastanum L. (Horse Chestnut)

through somatic embryogenesis. J Plant

Physiol 132:322–326

15. Chalupa V (1990) Somatic embryogenesis and

plant regeneration in Quercus petraea (Matt.)

Liebl., Tilia platyphyllos Scop., and Aesculus hip-

pocastanum L. Lestnictvi. Forestry 36:599–604

16. Capuana M, Debergh PC (1997) Improvement

of the maturation and germination of horse

chestnut somatic embryos. Plant Cell Tiss Org

Cult 48:23–29

17. Troch V, Werbrouck S, Geelen D, Van Labeke

M-C (2009) Optimization of horse chestnut

( Aesculus hippocastanum L.) somatic embryo

conversion. Plant Cell Tiss Org Cult 98:

115–123

Maurizio Capuana

439

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 24

Somatic Embryogenesis in Araucaria angustifolia

(Bertol.) Kuntze (Araucariaceae)

Miguel P. Guerra , Neusa Steiner , Francine L. Farias-Soares ,

Leila do N. Vieira , Hugo P.F. Fraga , Gladys D. Rogge-Renner ,

and Sara B. Maldonado

Abstract

This chapter deals with the features of somatic embryogenesis (SE) in Araucaria angustifolia , an

endangered and native conifer from south Brazil. In this species SE includes the induction and proliferation

of embryogenic cultures composed of pro-embryogenic masses (PEMs), which precede somatic embryos

development. A. angustifolia SE model encompasses induction, proliferation, pre-maturation, and

maturation steps. Double-staining with acetocarmine and Evan’s blue is useful to evaluate the embryonic

somatic structures. In this chapter we describe A. angustifolia SE protocols and analyzes morphological

features in the different SE developmental stages.

Key words Conifers , Forest biotechnology , Germplasm conservation , Plant cell culture , Plant

physiology , Somatic embryo genesis

1 Introduction

The Brazilian pine Araucaria angustifolia (Bertol.) Kuntze

( Araucariaceae ) is a native conifer with relevant economic impor-

tance in Brazil, representing the most exploited timber source until

the 1970s [

1 ]. Uncontrolled exploitation of the high-quality wood

has led to the species classiﬁ cation as critically endangered in the

International Union for the Conservation of Nature and Natural

Resources Red Book [

2 ]. In the last years, it has been suggested

for A. angustifolia conservation integrated ex situ and in situ strat-

egies to conserve genetic resources [

3 ]. In addition, the mainte-

nance of ex situ seed banks is not feasible for recalcitrant seeds,

such as A. angustifolia requiring the use of in vitro techniques to

germplasm conservation [

4 ].

Biotechnological tools have a large potential in breeding

and biodiversity conservation programs for woody species [

5 ].

440

In this sense, somatic embryo genesis (SE) has been successfully

applied for somatic cells and viable embryos obtaining, in a

morphogenetic process closely related to the natural process of

zygotic embryo genesis (ZE) [

6 ]. SE in A. angustifolia is a com-

plex and multifactorial pathway that includes induction and

proliferation of embryogenic cultures (EC), composed of pro-

embryogenic masses ( PEMs ) preceding somatic embryo forma-

tion [

1 , 5 , 7 ]. A. angustifolia SE model encompasses two cycles.

The cycle A consists in induction, proliferation and pre-matura-

tion steps. Induction is characterized by EC formation in zygotic

embryo apex (Fig.

1a ), which is retrieved and in vitro cultivated

in both auxin and cytokinin presence (Fig.

1b ) or in plant

growth regulator ( PGR )-free culture medium [

8 – 10 ]. Through

double-staining analysis with acetocarmine and Evan’s blue, it

is possible to identify in the PEMs the presence of two typical

conifer cells: embryogenic cells and suspensor -like cell s ( SCs )

[

11 – 13 ]. During proliferation step, PEMs evolve through

three specific developmental stages, PEM I, II, and III, evalu-

ated by the abundance of embryogenic cells and SCs [

10 – 12 ].

Fig. 1 Morphological aspects of Araucaria angustifolia embryogenic cultures. ( a ) Embryogenic callus 30 days after

somatic embryo genesis induction. ( b ) Embryogenic callus during multiplication cycles in gelled culture

medium. ( c ) Callus with globular-staged somatic embryos during maturation cycle. ( d ) Torpedo-staged somatic

embryo after 90 days in maturation culture medium ( arrows indicate globular-staged somatic embryos). Bar, 2 mm

Miguel P. Guerra et al.

441

PEMs-to-early somatic embryo transition is a central event in

conifers SE [

6 ]. In A. angustifolia SE, pre-maturation step is

the starting point of the early SE polarization and individualiza-

tion from PEM III [

7 ]. The trigger for this process is the PGR

removal of culture medium, followed by maltose and PEG sup-

plementation [

11 , 14 ]. Early somatic embryos arise when com-

pact clusters of embryogenic cells grow from PEM III with two

regions, the dense globular embryonal mass ( EM ) in the apical

part, and suspensor (S) in the basal part [

5 ]. After pre-matura-

tion step, in the cycle B, starts the maturation phase, where

early somatic embryos (Fig.

1c ) are able to develop in late

somatic embryos (Fig.

1d ). Late somatic embryos formation

can be achieved when the early embryos are capable to respond

to the new speciﬁ c signals with osmotic and hormonal adjust-

ment during maturation step [

3 , 7 , 11 ]. The initiation of early

somatic embryo formation can be observed with the embryonic

cell group increase, while the elongated suspensor cells undergo

programmed cell death [

9 , 13 – 15 ]. The early somatic embryo

development marks the beginning of structural differentiation

with the protoderm formation around the early somatic embryo

followed by the meristem determination (root and shoot apical

meristems). After that, the somatic embryos obtained can be

converted into plantlets. Thus, the approach of this chapter is to

describe SE protocols and describe morphological features of

SE developmental stages in A. angustifolia .

2 Materials

Prepare all solutions using distilled water and analytical grade

reagents. Prepare all stock solutions at room temperature. All stock

solutions can be autoclaved excepting solutions containing vita-

mins and amino acids.

1. Immature zygotic embryo s of A. angustifolia excised of seeds

collected from female cones in December.

2. 70 % (v/v) ethanol.

3. 2 % sodium hypochlorite.

4. Sterile distilled water.

5. Glass ﬂ asks.

1. BM-macrosalt solution [

16 ], 20×: Add about 500 mL of

distilled water to a 1000 mL glass beaker. Weigh 12.07 g

NH

4 NO

3 , 18.20 g KNO

3 , 2.72 g KH

2 PO

4 , 4.93 g

MgSO

4 ·7H

2 O, 5.13 g Mg(NO

3 )

2 ·4H

2 O, 1 g MgCl

2 ·6H

2 O,

and 4.72 g Ca(NO

3 )

2 ·4H

2 O, transfer to the beaker, and solu-

bilize. Make up to 1000 mL with water. Store at 4 °C.

2.1 Plant Material

and Surface

Sterilization

2.2 Stock Solutions

of the Induction

and Proliferation

Culture Medium

Somatic Embyogenesis in Araucaria angustifolia

442

2. BM-microsalt solution 200× [ 16 ]: Add about 500 mL of dis-

tilled water to a 1000 mL glass beaker. Weigh 1.59 g

MnSO

4 ·H

2 O, 2.82 g ZnSO

4 ·7H

2 O, 3.1 g H

3 BO

3 , 0.83 g Kl,

25 mg CuSO

4 ·H

2 O, 25 mg Na2MoO

4 ·5H2O, and 25 mg

CoCl

2 ·6H

2 O. Transfer to glass beaker, and solubilize. Make up

to 1000 mL with water. Store at 4 °C.

3. BM-amino acid solution 100×: Add 5 mL of distilled water to

a 10 mL glass beaker. Weigh 1 g

L -glutamine , 0.5 g casein, 1 g

myoinositol , transfer to beaker and solubilize. Make up to

10 mL with water. Prepare just before use, do not stock.

4. Fe-EDTA solution 20×: Add about 500 mL of distilled water

to a 1000 mL glass beaker. Weigh 187.2 mg Na

2 EDTA × 2H 2 O

and 139 mg FeSO

4 × 7H 2 O, transfer to beaker and solubilize.

Make up to 1000 mL with water. Store at 4 °C.

5. Vitamins and glycine solution 500×: Add about 500 mL of

distilled water to a 1000 mL glass beaker. Weigh 500 mg thia-

mine HCl, 250 mg pyridoxine HCl, 250 mg nicotinic acid , 1 g

glycine, add to the beaker and solubilize. Make up to 1000 mL

with water. Store aliquots of 2 mL microtubes at −20 °C.

1. MSG-macrosalt solution [

17 ], 20×: Add about 500 mL of dis-

tilled water to a 1000 mL glass beaker. Weigh 29 g NH

4 NO

3 ,

38 g KNO

3 , 8.8 g CaCl

2 ·2H

2 O, 3.4 g KH

2 PO

4 , 7.4 g

MgSO

4 ·7H

2 O, and 14.9 g KCl. Transfer to the beaker and

solubilize. Make up to 1000 mL with water. Store at 4 °C.

2. MSG-microsalt solution [

17 ], 200×: Add about 500 mL of

distilled water to a 1000 mL glass beaker. Weigh 3.38 g

MnSO

4 ·H

2 O, 1.72 g ZnSO

4 ·7H

2 O, 1.24 g H

3 BO

3 , 0.16 g Kl,

5 mg CuSO

4 ·H

2 O, 50 mg Na

2 MoO

4 ·5H

2 O, and 5 mg

CoCl

2 × 6H 2 O, transfer to the beaker, and solubilize. Make up

to 1000 mL with water. Store at 4 °C.

3. MSG-amino acid solution 100×: Add about 5 mL of distilled

water to a 10 mL glass beaker. Weigh 1.46 g

L -glutamine ,

0.1 g myoinositol , and transfer to the beaker and solubilize.

Make up to 10 mL with water. Prepare just before use, do not

stock.

4. Fe-EDTA solution 20×: Use the same solution described in

Subheading

2.2 .

5. Vitamins and glycine solution 500×: Use the same solution

described in Subheading

2.2 .

1. 1000 μM 2,4-Dichlorophenoxyacetic acid (2,4-D): Weigh

22.10 mg of 2,4-D and transfer to a 100 mL glass beaker. Add

1 mL of NaOH 1 M to dissolve 2,4-D. Make up to 100 mL

with water. Store at 4 °C ( see Note 1 ).

2.3 Stock Solutions

of the Pre- maturation

and Maturation

Culture Medium

2.4 Other Stock

Solutions

Miguel P. Guerra et al.

443

2. 1000 μM 6-benzylaminopurine (BAP): Weigh 22.50 mg of

BAP and transfer to a 100 mL glass beaker. Add 1 mL of

NaOH 1 M to dissolve BAP. Make up to 100 mL with water.

Store at 4 °C ( see Note 1 ).

3. 1000 μM kinetin (KIN): Weigh 21.50 mg of KIN and transfer to

a 100 mL glass beaker. Add 1 mL of NaOH 1 M to dissolve

KIN. Make up to 100 mL with water. Store at 4 °C ( see Note 1 ).

1. Sucrose .

2. Maltose .

3. Phytagel

® .

4. Gelrite

® .

5. Polyethylene glycol 3350.

6. Polyethylene glycol 4000.

7. Reduced

L -glutathione.

8. Abscisic acid ( ABA ).

9. Activated charcoal .

1. To prepare 1 L of BMi add 30 g of sucrose to 400 mL of water

in a 1000 mL glass beaker and stir on a magnetic stirrer. Add

50 mL of BM-macrosalt stock solution, 5 mL of BM-microsalt

stock solution, 5 mL of Fe-EDTA stock solution, 5 mL of

2,4-D stock solution, 2 mL of BAP stock solution, and 2 mL

of KIN stock solution.

2. At this step, PGR -free culture medium is also used for SE

induction.

3. Add water to just under the ﬁ nal volume of 988 mL. While

stirring, adjust the pH by adding 0.5 M NaOH or 0.5 M HCl

solution to reach a pH of 5.8 and add 2 g of Phytagel

® .

Autoclave for 15 min at 121 °C.

4. Wait the autoclaved mixture temperature to reach 40 °C. In

the laminar air ﬂ ow cabinet, add the ﬁ lter-sterilized ( see Note 3 )

solution containing 10 mL of BM-amino acid stock solution

and 2 mL of vitamins and glycine stock solution. Adjust pH by

adding 0.5 M NaOH or 0.5 M HCl solution to reach a pH of

5.8 before ﬁ lter-sterilization.

1. To prepare 1 L of BMp add 30 g of sucrose to 400 mL of water

in a 1000 mL glass beaker and stir on a magnetic stirrer. Add

50 mL of BM-macrosalt stock solution, 5 mL of BM-microsalt

stock solution, 5 mL of Fe-EDTA stock solution, 2 mL of

2,4-D stock solution, 0.5 mL of BAP stock solution, and

0.5 mL of KIN stock solution.

2.5 Culture Medium

Supplements

2.6 Culture

Medium Preparation

( See Note 2 )

2.6.1 Induction Culture

Medium (BMi)

2.6.2 Proliferation

Culture Medium (BMp)

Somatic Embyogenesis in Araucaria angustifolia

444

2. Cultures induced in PGR -free culture medium can be multiplied

either in the culture medium described in 1, or in PGR- free

culture medium.

3. Add water to just under the ﬁ nal volume of 988 mL. While

stirring, adjust the pH by adding 0.5 M NaOH or 0.5 M HCl

solution to reach a pH of 5.8 and add 2 g of Phytagel

® .

Alternatively the EC can be multiplied in liquid medium.

Autoclave for 15 min at 121 °C.

4. Wait the autoclaved mixture temperature to reach 40 °C. In

the laminar airﬂ ow cabinet, add the ﬁ lter-sterilized ( see Note

3 ) solution containing 10 mL of BM-amino acid stock solu-

tion and 2 mL of vitamin and glycine stock solution. Adjust

pH by adding 0.5 M NaOH or 0.5 M HCl solution to reach a

pH of 5.8 before ﬁ lter-sterilization.

5. For gelled culture medium, shake the solution to homogenize

the mixture while warming. Distribute the mixture by pouring

into sterile 15 mm × 90 mm Petri dishes (1 L of culture medium

provides ~40 dishes). Leave the dishes to cool and solidify.

Close and seal the dishes with Paraﬁ lm

® .

6. For liquid culture medium, shake the solution to homogenize

the mixture. Distribute 50 mL of the mixture into a sterile

250 mL Erlenmeyer ﬂ ask. Close and seal with Paraﬁ lm

® .

Recently, two pre-maturation protocols have been described for

A. angustifolia SE and both of them can be successfully applied

[

14 , 18 ].

1. To prepare 500 mL of pre-maturation culture medium

(MSGpm1), add 45 g of maltose and 35 g of PEG 3350–

200 mL of water in a 500 mL glass beaker and stir on a mag-

netic stirrer. Add 25 mL of MSG-macrosalt stock solution,

2.5 mL of MSG-microsalt stock solution, 2.5 mL of Fe-EDTA

stock solution.

2. Add water to just under the ﬁ nal volume of 494 mL. While

stirring, adjust the pH by adding 0.5 M NaOH or 0.5 M HCl

solution to reach a pH of 5.8. Autoclave for 15 min at 121 °C

in a 750–1000 mL Erlenmeyer ﬂ ask.

3. Wait the temperature to reach 40 °C and add the ﬁ lter-steril-

ized ( see Note 3 ) solution containing 5 mL of MSG-amino

acid stock solution, 1 mL of vitamin and glycine stock solu-

tion, and 1.53 g of reduced

L -glutathione in the laminar air-

ﬂ ow cabinet. Adjust pH by adding 0.5 M NaOH or 0.5 M

HCl solution to reach a pH of 5.8 before ﬁ lter-sterilization.

This procedure should be done preferably in the dark to pre-

vent reduced

L - glutathione degradation.

2.6.3 Pre-maturation

Culture Medium 1 [ 14 ]

Miguel P. Guerra et al.

445

4. Shake the solution to homogenize the mixture. Distribute

2 mL of the mixture by pipetting into sterile 12-well culture

plate (500 mL of culture medium provides ~20 multiwell cul-

ture plates). Close and seal with Paraﬁ lm

® .

1. To prepare 500 mL of pre-maturation culture medium

(MSGpm2), add 15 g of sucrose , 35 g of maltose , 45 g of PEG

4000, and 1.5 g of activated charcoal to 200 mL of water in a

500 mL glass beaker and stir on a magnetic stirrer. Add 25 mL

of MSG-macrosalt stock solution, 2.5 mL of MSG-microsalt

stock solution, and 2.5 mL of Fe-EDTA stock solution.

2. Add water to just under the ﬁ nal volume of 494 mL. While

stirring, adjust the pH by adding 0.5 M NaOH or 0.5 M

HCl solution to reach a pH of 5.7. Add 1.5 g of Gelrite

®

and autoclave for 15 min at 121 °C in a 750–1000 mL

Erlenmeyer ﬂ ask.

3. In the laminar airﬂ ow cabinet, wait the temperature to reach

40 °C and add the ﬁ lter–sterilized ( see Note 3 ) solution con-

taining 0.73 g of

L -glutamine , and 1 mL of vitamin and glycine

stock solution. Adjust pH by adding 0.5 M NaOH or 0.5 M

HCl solution to reach a pH of 5.7 before ﬁ lter-sterilization.

4. Shake the solution to homogenize the mixture while warm.

Distribute the mixture by pouring into sterile 15 mm × 90 mm

Petri dishes (500 mL of culture medium provides ~20 dishes).

Leave the dishes to cool and solidify. Close and seal the dishes

with Paraﬁ lm

® .

1. To prepare 1 L of maturation culture medium (BMm), add

90 g of maltose , 70 g of PEG 3350 and 1.5 g of activated char-

coal to 400 mL of water in a 1000 mL glass beaker and stir on

a magnetic stirrer. Add 50 mL of BM-macrosalt stock solution,

5 mL of BM-microsalt stock solution, and 5 mL of Fe-EDTA

stock solution.

2. Add water to just under the ﬁ nal volume of 988 mL. While

stirring, adjust the pH by adding 0.5 M NaOH or 0.5 M HCl

solution to reach a pH of 5.8 and add 2 g of Phytagel

® .

Autoclave for 15 min at 121 °C.

3. Wait the temperature to reach 40 °C and add the ﬁ lter-steril-

ized ( see Note 3 ) solution containing 31.7 mg of ABA ( see

Note 4 ), 10 mL of BM-amino acid stock solution, and 2 mL

of vitamin and glycine stock solution in the laminar ﬂ ow cabi-

net. Adjust the solution pH by adding 0.5 M NaOH or 0.5 M

HCl solution to reach a pH of 5.8 before ﬁ lter-sterilization.

4. Shake the solution to homogenize the mixture while warm.

Distribute the mixture by pouring into sterile 15 mm × 90 mm

Petri dishes (1 L of culture medium provides ~40 dishes).

Leave the dishes to cool and solidify. Close and seal the dishes

with Paraﬁ lm

® .

2.6.4 Pre-maturation

Culture Medium 2 [ 18 ]

2.6.5 Maturation

Culture Medium

Somatic Embyogenesis in Araucaria angustifolia

446

1. 2 % carmine: Dissolve 2 g carmine in 100 mL acetic acid 45 %

(v/v). Boil in reﬂ ux condenser for 3 h. Cool at room tempera-

ture and ﬁ lter with ﬁ lter paper.

2. 0.05 % (w/v) Evan’s blue: Dissolve 1 g Evan’s blue in 100 mL

distilled water.

3. Slides and cover glass.

4. Light microscope.

1. Scalpel, forceps.

2. Spirit burner.

3. Magnetic stirrer.

4. pH meter, autoclave.

5. Bottles, Petri dishes, Erlenmeyer ﬂ asks, glass beaker, 12-well

culture plate.

6. Paraﬁ lm

® .

7. Syringe, sterile syringe ﬁ lters Chromaﬁ l

® , ﬁ lter paper.

8. Analytical balance.

9. 0.5 N sodium hydroxide (NaOH), 0.5 N hydrochloric acid

(HCl).

10. Incubator chamber, laminar ﬂ ow cabinet, stereomicroscope.

3 Methods

All the procedures described below must be performed in laminar

ﬂ ow cabinet, with sterilized instruments.

1. Only immature seeds of A. angustifolia with globular-staged

zygotic embryo s are used, in order to induce SE. Surface steril-

ize seeds in a glass beaker with 70 % ethanol for 5 min. Remove

ethanol and add 2 % sodium hypochlorite for 20 min. Remove

sodium hypochlorite and wash seeds three times with auto-

claved distilled water. All solutions must be added in enough

volume to cover the seeds into the beaker.

2. With the aid of a stereomicroscope, scalpel, and forceps on a

sterilized Petri dish, excise the immature zygotic embryo and

inoculate into the induction culture medium. Cultures are

maintained in BOD incubator chamber at 24 ± 2 °C.

After 30-day culture in BMi culture medium, EC is generally

obtained. During proliferation step, EC are composed by PEMs ,

maintained in repetitive multiplication cycles for an undetermined

period of time. At this point, EC proliferation can be achieved with

or without PGR supplementation. Proliferation can also be per-

formed in gelled or liquid culture medium.

2.7 Cytochemical

Analysis

2.8 Other Useful

Materials

3.1 SE Induction

3.2 EC Proliferation

Miguel P. Guerra et al.

447

1. To perform the subculture for gelled BMp culture medium,

friable and translucent EC should be removed from the BMi

medium with the aid of a forceps and transferred to fresh gelled

BMp culture medium. Colonies of cells should be mixed dur-

ing the process of subculture to promote uniform distribution

of nutrients contained in culture medium. The subculture pro-

cedure must be performed every 21 days to fresh gelled BMp

culture medium and can be done indeﬁ nitely. Cultures are

maintained in BOD incubator chamber at 24 ± 2 °C.

2. For liquid BMp culture medium, about 500 mg of friable and

translucent EC should be removed from the BMi medium

with the aid of a forceps and transferred to a fresh liquid BMp

medium. The subculture procedure must be performed every

15 days to a fresh liquid BMp culture medium and can be

done indeﬁ nitely. This procedure is realized with the aid of

“Cell Dissociation Sieve” (Sigma-Aldrich), 80 mesh screens.

Capture the EC by pouring the culture medium with EC in

proliferation in the “Cell Dissociation Sieve.” With the aid of

a forceps, take 500 mg of EC and transfer to a new ﬂ ask.

Cultures are maintained in an orbital shaker at 90 rpm, at

24 ± 2 °C in the dark.

Pre-maturation is an important step in conifers SE, and it was

recently applied to A. angustifolia protocol [

14 , 18 ]. In this step,

the transition of PEMs to early somatic embryo s is observed.

1. After proliferation step, repeat the same procedure described

above ( see Subheading

3.2 ) to capture the EC. Transfer about

50 mg of EC with the aid of a forceps to a 12-well culture plate

containing 2 mL MSGpm1 per well.

2. The plates should be incubated in an orbital shaker at 90 rpm

in the dark. Cultures are maintained in a growth room at

24 ± 2 °C for 15 days.

1. After proliferation step, repeat the same procedure described

above (Subheading

3.2 , step 2 ) to capture the EC. Transfer

100 mg of EC with the aid of a forceps to a sterile ﬁ lter paper

disc (Ø 80 mm). Transfer the ﬁ lter paper with the cultures to

Petri dish containing MSGpm2 culture medium.

2. Cultures are maintained in a growth room at 24 ± 2 °C for 30

days.

1. For somatic embryo s maturation, about 500 mg of EC con-

taining early somatic embryos is transferred with the aid of a

forceps to BMm culture medium.

2. Petri dishes are maintained in BOD incubator chamber at

24 ± 2 °C for 60 days. One subculture should be performed at

day 30 in culture to a fresh BMm culture medium.

3.3 EC Pre-

maturation 1

3.4 EC Pre-

maturation 2 [ 18 ]

3.5 Early Somatic

Embryo Maturation

Somatic Embyogenesis in Araucaria angustifolia

448

The quality of cultures is evaluated by double staining under light

microscope based on acetocarmine and Evan’s blue staining [

19 ].

This double-staining analysis reveals the presence of the two typical

embryonic conifer structures: the embryogenic cells, which are iso-

diametric and densely cytoplasmic, reacting in red with acetocar-

mine, and the suspensor -like cell s , which are vacuolated and reacts

in blue to Evan’s blue [

20 ].

1. Take an aliquot of 50 mg of EC and transfer to a watch glass.

2. Add a drop of 1 % acetocarmine (w/v) to the sample, gently

mix, and wait for 1 min.

3. Carefully remove the acetocarmine with the aid of toilet paper.

4. Drop 0.05 % Evan’s blue (w/v) to the sample, gently mix and

wait 1 min.

5. Carefully remove the Evan’s blue with the aid of toilet paper.

6. Drop 1 mL of sterile distilled water.

3.6 Morphological

and Cytochemical

Analysis Procedure

Fig. 2 Araucaria angustifolia embryogenic cultures morphological and cytochemical analysis with acetocar-

mine and Evan’s blue. ( a – c ) Proembryogenic masses at PEM I stage ( a ), PEM II stage ( b ) and PEM III stage ( c ).

( d ) PEM III-staged embryogenic cells starting polarization and individualization process. ( e ) Early somatic

embryo s (ESE) individualized and polarized. ( f ) Globular-staged ESE. EC, embryogenic cells stained with ace-

tocarmine; SC , suspensor -like cell s stained with Evan’s blue

Miguel P. Guerra et al.

449

7. Drop with a pipette an aliquot on a slide glass, and then visual-

ize in the light microscope.

8. Analyze and quantify the presence of PEM I (Fig.

2a ), PEM II

(Fig.

2b ), PEM III (Fig. 2c ), and early somatic embryo s

(Fig.

2e, f ) as well as the presence of SCs (Fig. 2a ) and embryo-

genic cells (Fig.

2a, b ).

4 Notes

1. The PGR stock solutions can be autoclaved for 15 min to

decrease bacterial and fungal contamination, and improve the

solubilization.

2. Culture medium should be prepared at least 3 days before the

inoculation procedure. This is the required period to ensure that

there was no fungal or bacterial contamination during the cul-

ture medium preparation.

3. Filter-sterilization is made with the aid of a syringe and ster-

ile Syringe ﬁ lters Chromaﬁ l

® (Macherey-Nagel), with PTFE

membrane, 0.20 μm pore size into the laminar ﬂ ow

cabinet.

4. Abscisic acid cannot be maintained in stock solution. Weigh

the abscisic acid with the aid of a analytical balance, add

200 μL of NaOH 1 M to dissolve ABA , and then add the

vitamins, amino acids, or other stock solutions you need to

ﬁ lter-sterilize.

References

1. Guerra MP, Steiner N, Mantovani A, Nodari

RO, Reis MS, Santos KL (2008) Araucária:

Evolução, ontogênese e diversidade genética.

In: Barbieri RL, Stumpf ERT (eds) Origem e

evolução de plantas cultivadas, 1st edn.

Embrapa Informação Tecnológica, Brasília,

pp 149–184

2. IUCN, IUCN Red List of Threatened Species,

Version 2011.2.

www.iucnredlist.org ,

Downloaded on 4.04.2012

3. Stefenon VM, Steiner N, Guerra MP, Nodari

RO (2009) Integrating approaches towards

the conservation of forest genetic resources: a

case study of Araucaria angustifolia . Biodivers

Conserv 18:2433–2448

4. Farias-Soares FL, Burrieza HP, Steiner N,

Maldonado S, Guerra MP (2013) Immunoanalysis

of dehydrins in Araucaria angustifolia embryos.

Protoplasma 250:911–918

5. Steiner N, Santa-Catarina C, Andrade JBR,

Balbuena TS, Guerra MP, Handro W, Floh

EIS, Silveira V (2008) Araucaria angustifolia

biotechnology-review. Funct Plant Sci

Biotechnol 2:20–28

6. von Arnold S, Sabala I, Bozkov P, Dyachok J,

Filonova L (2002) Developmental pathways of

somatic embryogenesis. Plant Cell Tiss Organ

Cult 69:233–249

7. Steiner N, Vieira FN, Maldonado S, Guerra

MP (2005) Carbon source affects morphogen-

esis and histodifferentiation of A. angustifolia

embryogenic cultures. Braz Arch Biol Technol

48:896–903

8. Astarita LV, Guerra MP (1998) Early somatic

embryogenesis in Araucaria angustifolia -

induction and maintenance of embryonal-

suspensor mass cultures. Braz J Plant Physiol

10:113–118

Somatic Embyogenesis in Araucaria angustifolia

450

9. Santos ALW, Silveira V, Steiner N, Vidor M,

Guerra MP (2002) Somatic embryogenesis in

Paraná Pine ( Araucaria angustifolia (Bert.)

O. Kuntze). Braz Arch Biol Technol

45:97–106

10. Silveira V, Steiner N, Santos ALW, Nodari RO,

Guerra MP (2002) Biotechnology tools in

Araucaria angustifolia conservation and

improvement: inductive factors affecting

somatic embryogenesis. Crop Breed Appl

Biotechnol 2:463–470

11. Santos ALW, Steiner N, Guerra MP, Zoglauer

K, Moerschbacher BM (2008) Somatic

embryogenesis in Araucaria angustifolia . Biol

Plant 52:95–99

12. Santos ALW, Silveira V, Steiner N, Maraschin

M, Guerra MP (2010) Biochemical and mor-

phological changes during the growth kinetics

of Araucaria angustifolia suspension cultures.

Braz Arch Biol Technol 53:497–504

13. Dutra NT, Silveira V, de Azevedo IG, Gomes-

Neto LR, Façanha AR, Steiner N, Guerra MP,

Floh EIS, Santa-Catarina C (2013) Polyamines

affect the cellular growth and structure of pro-

embryogenic masses in Araucaria angustifolia

embryogenic cultures through the modulation

of proton pump activities and endogenous lev-

els of polyamines. Physiol Plant 148:121–132

14. Vieira LN, Santa-Catarina C, Fraga HPF, dos

Santos ALW, Steinmacher DA, Schlogl PS,

Silveira V, Steiner N, Floh EIS, Guerra MP

(2012) Glutathione improves early somatic

embryogenesis in Araucaria angustifolia

(Bert) O. Kuntze by alteration in nitric oxide

emission. Plant Sci 195:80–87

15. Silveira V, Santa-Catarina C, Tun NN, Scherer

GFE, Handro W, Guerra MP, Floh EIS (2006)

Polyamine effects on the endogenous poly-

amine contents, nitric oxide release, growth

and differentiation of embryogenic suspension

cultures of Araucaria angustifolia (Bert.)

O. Kuntze. Plant Sci 171:91–98

16. Gupta PK, Pullman GS (1991) Method for

reproducing coniferous plants by somatic

embryogenesis using abscisic acid and osmotic

potential variation, US patent 5, 36–37

17. Becwar MR, Wann SR, Johnson MA, Verhagen

SA, Feirer RP, Nagmani R (1988) Development

and characterization of in vitro embryogenic

systems in conifers. In: Ahuja MR (ed) Somatic

cell genetics of woody plants. Kluwer

Academic, Dordrecht, pp 1–18

18. Jo L, dos Santos ALW, Bueno CA, Barbosa HR,

Floh EIS (2013) Proteomic analysis and poly-

amines, ethylene and reactive oxygen species lev-

els of Araucaria angustifolia (Brazilian pine)

embryogenic cultures with different embryo-

genic potential. Tree Physiol 34:94–104

19.

Gupta PK, Durzan DJ (1987) Biotechnology

of somatic polyembryogenesis and plantlet

regeneration in loblolly pine. Nat Biotechnol

5:147–151

20. Filonova LH, Bozhkov PV, von Arnold S

(2000) Developmental pathway of somatic

embryogenesis in Picea abies as revealed by

time-lapse tracking. J Exp Bot 51:249–264

Miguel P. Guerra et al.

Part IV

Protocols of Gametic Embryogenesis

in Selected Higher Plants

453

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 25

Anther Culture in Eggplant ( Solanum melongena L.)

Giuseppe Leonardo Rotino

Abstract

The technique of in vitro anther culture is the most favorite to incite the production of plants from micro-

spore through direct embryogenesis or regeneration from callus. Anther culture has been employed since

1980s in eggplant to obtain double-haploid plants from microspore derived embryos. From that time it

has been reﬁ ned and widely applied both at commercial level for a fast generation double-haploid parental

lines of F1 hybrids, as well as for experimental studies as the complete homozygosis of the microspore-

derived plants make more simply the genetic analysis. In this chapter, a step-by-step procedure is reported,

taking into consideration all the aspects of the technique, including the growth condition of the anther

donor plant, the in vitro regeneration of the androgenetic plantlets, their ploidy analysis, and the colchi-

cine treatment to double the chromosome number of the haploids.

Key words Androgenesis , Double- haploid , Haploid , Tissue culture , Microspore

1 Introduction

Haploidy through natural parthenogenesis has never been obser ved

in eggplant . The ﬁ rst haploids in eggplant were obtained by the

“Chinese Research Group of Haploid Breeding” [

1 ] and by

Isouard et al. [

2 ]. Then, Dumas de Vaulx and Chambonnet [ 3 ]

and Chambonnet [

4 ] greatly improved the yield of in vitro anther

derived plantlets by using a method similar to the one applied to

pepper [

5 ]. This protocol is based on a high temperature (35 °C)

treatment during the ﬁ rst period of anther culture . Studies on egg-

plant isolated- microspore culture have been carried out and plant-

lets were regenerated from microspore-derived callus following

either anther pre-culture [

6 ] or direct culture of microspores [ 7 ,

8 ]. However, no consistent technique has been so far published

about a direct regeneration of plantlets at a satisfactory rate through

embryogenesis from isolated microspore culture.

454

Dumas de Vaulx and Chambonnet method [ 3 ] resulted in a

reliable protocol to produce pollen -derived plants, enabling a

successful integration of doubled haploid lines in eggplant breed-

ing programs. Subsequent minor modiﬁ cations of this method

allowed to further enlarge the usefulness of in vitro androgenesis

for the production of modern eggplant varieties [

9 ]. According to

this method, excised anthers are cultured in the induction medium

(C), supplemented with appropriate plant growth regulator s

( PGR ), and placed in the darkness at 35 °C for 8 days of culture.

In the following days, Petri dishes are kept in the growth chamber

at 25 °C under a 16-h illumination (50 μmol/m

2 /s, ﬂ uorescent

light). On the 13th day, anthers are transferred to the differentia-

tion medium (R). Generally, embryos become visible from the

anthers after 1 month from the beginning of culture and the

embryo production lasts for 3–4 months. Well-formed embryos of

4–6 mm are transferred to the PGR-free medium (V3) for further

development. Complete plantlets can easily be propagated in vitro

by cuttings, using the apical bud with 3–4 nodes, and transferred

to soil. Hereafter, a slightly modiﬁ ed Dumas de Vaulx protocol

[

3 ], regarding the PGR composition of the culture media, is

described, together with advices on the various steps of the proce-

dure. Pluriannual observations about the response of different

genotypes evidenced that some of them were able to regenerate

androgenetic plants only in a medium containing speciﬁ c PGR

combinations. For this reason, it is suggested to try simultaneously

alternative induction and regeneration media, especially for novel

donor material or segregating progenies (Table

1 ).

Table 1

Sucrose content and PGR composition of the induction (C3, C6, C9, and

C13) and regeneration media (R1K and R1Z) media for anther culture of

eggplant (plus 8 g/L agar ; pH 5.9 before autoclaving. KIN, kinetin; 2,4-D,

2,4-dichlorophenoxyacetic acid; IAA, indole-3-acetic acid; NAA,

1-naphthaleneacetic acid; ZEA, zeatin)

Compounds

Induction (C) Regeneration (R)

C3 C6 C9 C13 R1K R1Z

Sucrose (g/L) 30 30 30 120 30 30

KIN (mg/L) 3 5 – 5 0.1 –

2,4-D (mg/L) – – – 5 – –

IAA (mg/L) 1 – – – – –

NAA (mg/L) – 5 3 – – –

ZEA (mg/L) – – 1 – – 0.1

Giuseppe Leonardo Rotino

455

2 Materials

The highly responsive cv Dourga can be employed to start practic-

ing the protocol, with plants preferably grown in greenhouse.

However, also donor plants from the open ﬁ eld can be used.

1. Laminar ﬂ ow hood.

2. Sterile Whatman or blotting paper.

3. Forceps, scalpels and spatulas.

4. Petri dishes, 60 mm diameter.

5. Glass or plastic jars (e.g., 250 and 500 mL, Magenta box).

6. Autoclave.

7. Plastic wrap ﬁ lm.

8. Stereo and optical ﬂ uorescence microscopes.

9. Microscope slides and cover slips.

10. Incubator at 35 °C.

11. Stainless steel tea mesh infuser spoon.

12. Growth chambers.

13. Flow cytometer.

1. Tween 20 and dish soap.

2. 80 % ethanol.

3.

Sodium hypochlorite.

4. Sterile deionized water.

5. TRIS.

6. Triton X-100.

7. Induction (C), regeneration (R), and multiplication (V3)

media (Tables

1 and 2 ).

8. Acetocarmine [1 g carmine, 100 mL glacial acetic acid (45 %),

5 mL FeCl · 6H

2 O].

9. Lysis solution for leaf mesophyll nuclei extraction.

10. Staining solution for nuclear DNA (Partec).

11. FDA stock solution: Fluorescein-diacetate dissolved into ace-

tone (5 mg/mL).

12. Colchicine .

13. Lanoline.

2.1 Plant Materials

2.2 Equipment

2.3 Solutions

and Media

Anther Culture in Eggplant

Table 2

Macronutrients, micronutrients, and vitamins of the three basal media

utilized for eggplant anther culture (mg/L)

Macroelements a Media

C R V3

KNO

3 2150 2150 1900

NH

4 NO

3 1238 1238 1650

MgSO

4 · 7H 2 O 412 412 370

CaCl

2 · 2H 2 O 313 313 440

KH

2 PO

4 142 142 170

Ca(NO

3 )

2 · 4H 2 O 50 50 –

NaH

2 PO

4 · H 2 O 38 38 –

(NH

4 )

2 SO

4 34 34 –

KCl 7 7 –

Microelements

a

MnSO

4 · H 2 O 22.130 20.130 0.076

ZnSO

4 · 7H 2 O 3.625 3.225 1.000

H

3 BO

3 3.150 1.550 1.000

KI 0.695 0.330 0.010

Na

2 MoO

4 · 2H 2 O 0.188 0.138 –

CuSO

4 · 5H 2 O 0.016 0.011 0.030

CoCl

2 · 6H 2 O 0.016 0.011 –

AlCl

3 · 6H 2 O – – 0.050

NiCl

2 · 6H 2 O – – 0.030

Vitamins and amino acids

Myoinositol 100.00 100.00 100.00

Pyridoxine HCl 5.500 5.500 5.500

Nicotinic acid 0.700 0.700 0.700

Thiamine HCl 0.600 0.600 0.600

Calcium pantothenate 0.500 0.500 0.500

Vitamin B12 0.030 – –

Biotin 0.005 0.005 0.005

Glycin 0.100 0.100 0.200

Chelated iron

Na

2 EDTA 18.65 18.65 37.30

FeSO

4 · 7H 2 O 13.90 13.90 27.80

a As reported in [ 4 ]

457

3 Methods

1. Generally, healthy and vigorous eggplants provide anthers with

the highest androgenetic potential. It is important to prevent

seed-setting and plant aging by removing open ﬂ owers and

small fruits. It is important to control insect and mite attacks;

however anthers should be collected several days after spraying

pesticide ( see Note 1 ).

2. Detach from the donor plants the ﬂ ower buds with, roughly,

the upper fused edge of the sepal, almost of the same height of

the petals (Fig.

1 , see Note 2 ). This stage of bud development

ensures that a large part of the microspores is at the uninucle-

ate or the very early binucleate stage of development which,

generally, gives better results. Such evidences have been

recently conﬁ rmed by Salas et al. [

10 ] , which demonstrated

that vacuolate microspores and young bicellular pollen are the

most responsive stages when microspores are directly cultured

in liquid medium, whereas cultured anthers, mostly containing

microspores at these stages, displayed a reduced or even null

androgenetic response. The authors ascribed such discrepancy

to a delayed contact of the media components with the micro-

spores which became too old and lose their androgenic induc-

ible state because of the time needed by the active substances

to reach the anther locule.

1. Collected ﬂ ower buds are brieﬂ y immersed and gently stirred

into a solution of demineralised water with a few drops of Tween

20 or dish soap, followed by 30 s in a solution of 80 % ethanol.

3.1 Anther Donor

Plants and Choice

of Floral Buds

3.2 Sterilization

and In Vitro Culture

Fig. 1 Flower buds at different developmental stages. The six buds in the middle are suitable for anther culture ,

whereas the two buds on the right and the two buds on the left are, respectively, too young and too old

Anther Culture in Eggplant

458

Then, in the laminar ﬂ ow hood, the ﬂ oral buds are immersed

for 20 min in a solution of 30 % commercial bleach (1 % active

chlorine), and ﬁ nally rinsed three times with sterile demineral-

ized water (3–4 min for each washing) (Fig.

2 , see Note 3 ).

2. The ﬂ ower buds are placed over a sterile paper to excise the

anthers by using a scalpel and a forceps (Figs.

3 and 4 ). Plate,

in a 60 mm Petri dish, 10–12 anthers with their concave (exter-

nal) zone onto medium C (Fig.

4d ). The Petri dishes are sealed

using household plastic wrap. Keep plated anthers at 35 °C in

the darkness for 8 days ( see Note 4 ).

Fig. 2 Buds sterilization. ( a ) Buds inside the steel tea mesh infusers. ( b ) Tea mesh containing the buds

immersed in the chlorine sterilization solution next to a Magenta box ﬁ lled with water for the ﬁ rst washing

Fig. 3 Extraction of the petal cone containing the anthers from the buds. ( a ) Transversal cut of bud below the

anther. ( b ) Longitudinal cut of the sepal. ( c ) Extraction of the petal cone containing the anthers

Giuseppe Leonardo Rotino

459

3. Transport and keep the anthers at 25 °C, 16-h photoperiod

(50 μmol/m

2 /s).

4. After 4 days transfer the anthers to R medium ( see Note 5 ).

5. Every 5–6 weeks transfer anthers to fresh R medium ( see Note 5 ).

6. Move the embryos sprouting out from the anthers to a 60 mm

Petri dish, containing either the V3 medium, if they have a well

formed principal root, or the R medium in the case the embryos

are younger (3–4 mm) ( see Note 6 ).

7. Well formed in vitro plantlets, with good root system and foli-

age, are acclimatized under growth chamber condition (16-h

light at 22 °C, 8-h dark at 18 °C, light intensity ~200 μmol/

m

2 /s) by transplanting into pots (Fig. 5 , see Note 7 ).

Fig. 4 Extraction and plating of the anthers. ( a ) Petal cone cut and opened. ( b ) Excision of anther by pushing to

its ﬁ lament. ( c ) Anther stuck with forceps. ( d ) Anthers plated in the C medium

Anther Culture in Eggplant

460

8. After 1–2 weeks, acclimatized plants can be moved to the

greenhouse, keeping them under shadow if the temperature

and light intensity are high.

Analysis of ploidy determination can be performed through direct

quantiﬁ cation of nuclear DNA content by ﬂ ow cytometer, or indi-

3.3 Ploidy

Determination

Fig. 5 From androgenetic embryos to plantlets. ( a ) An anther culture d in the R medium giving rise to several

microspore -derived embryos at different developmental stages. ( b ) Cultured anthers producing embryos or

callus. ( c ) From left to right : rooted androgenetic plants in the V3 medium ready to be transferred to soil, freshly

plantlet in ex vitro condition, and acclimatized plantlets, ready to be transferred to the greenhouse

Giuseppe Leonardo Rotino

461

rectly by counting the number of chloroplasts in the stomata guard

cells (Fig.

6a ). Leaves from in vitro rooted plantlets are the best

material for both the analyses ( see Note 8 ).

1. About 0.5 cm

2 of a young leaf of an in vitro grown plant is

chopped in the extraction buffer in a small Petri dish.

2. Filter (30 μm mesh) to enrich the solution with the nuclei; add

the staining buffer containing the ﬂ uorophore ( DAPI ,

Fluorescein, etc.) and keep at 4 °C for 1 h.

3. Start the analysis.

3.3.1 Flow Cytometry

Analysis According to the

Manufacturer Protocol

(E.g., Partec CyFlow, SL)

( see Note 9 )

Fig. 6 Ploidy determination and colchicine treatment. ( a ) Flow cytometric determination of DNA content in the

nuclei and number of chloroplasts in the stomata guard cells of a diploid and haploid androgenetic plant,

obtained from the same anther donor. ( b ) Lanoline paste containing 0.5 % colchicine, applied to the secondary

axillary buds of a haploid plant

Anther Culture in Eggplant

462

1. Cut a piece of 0.3–0.5 cm

2 of leaf and put it on the microscope

slide with the lower leaf lamina upward.

2. Wet the lower epidermal with 1–2 drops of FDA stock solution

diluted in water, add the cover slip, and wait for a maximum of

20–30 s.

3. Observe under a ﬂ uorescent light. The chloroplast will appear

green-colored in the stomata cells ( see Note 10 ).

Haploid plants, apart specific employment (e.g., basic research),

need to be treated to restore their diploid status, so that selfed

seeds can be obtained and the double- haploid lines established.

Chromosome doubling may be performed in several ways. Here it

is described the treatment of the secondary axillary buds with col-

chicine in the already acclimatized plantlets which, generally, ensure

that 50–70 % of the treated plants will produce seeds after selfing.

1. Use preferably fast growing plantlets with 4–8 leaves. Trim the

apical bud (to suppress apical dominance) and remove the axil-

lary buds with a scalpel.

2. Dissolve colchicine (0.5 %) in lanoline paste.

3. Apply with the aid of a spatula the lanoline-containing colchi-

cine to the secondary axillary buds.

4. Keep the treated plantlets in the dark for 48 h and then transfer

them back to greenhouse conditions.

5. Remove the shoots produced by the untreated buds (e.g.,

those below the soil level) (Fig.

6b , see Note 11 ).

4 Notes

1. Avoid excessive fruit setting, especially the presence of mature

and overripe fruits. Unfortunately, it is not known the environ-

mental conditions that allow to maximize the androgenetic

response of the anther donor eggplant . Seasonal variation has

been observed in the yield of eggplant androgenetic embryos.

It has been reported [

11 ] that during the period July–October

(in the Northern Hemisphere), the highest number of respond-

ing anthers were found from the middle of September until the

middle of October. These results are in accordance with our

observations that, in the Mediterranean climate, higher andro-

genetic frequencies are obtained during cooler months, and

the best periods are spring and autumn (unpublished). Most

likely, the photoperiod, as well as the day/night temperature,

affects anther response. More precise information could be

obtained by growing donor plants in a phytotron.

2. For beginners, it is suggested to check the exact stage of micro-

spore development during the various step of the ﬂ ower bud

3.3.2 Chloroplast

Counting in Stomata Guard

Cells ( See Note 10 )

3.4 Diploidization of

the Haploid Plants

Giuseppe Leonardo Rotino

463

growth. This is important because the sepal coverage of the

petal strongly varies among the different eggplant genotypes; it

is inﬂ uenced by plant ageing and is also affected by the environ-

ment. Anyway, due to this appreciable variation, it is suggested

to collect also ﬂ ower buds either slightly larger, or slightly

smaller than the ones considered of optimal size. For the cyto-

logical analysis of the microspore stage: squash one anther on

freshly prepared DAPI -TRIS buffer (TRIS buffer: 0.05 M

Tris–HCl, 0.5 % Triton X-100, 5 % sucrose , pH 7) and observe

under UV light. Alternatively, the anther can be squashed in

acetocarmine and observed under optical microscope.

3. The use of stainless steel spoons (like the ones used as tea mesh

infuser) makes very easy the sterilization of ﬂ ower buds, as it is

only necessary to move the spoon from one jar to the next

(Fig.

2 ). This ensures that the whole ﬂ owers are completely

immersed in the sterilization and washing solutions. After

washing, the buds are kept into a Petri dish, or left in the in tea

mesh infuser within the empty jar.

4. Extract the anthers by cutting transversally the ﬂ ower bud at its

base, in correspondence of the anther ﬁ lament or slightly

below (Fig.

3a ). Then, cut longitudinally only the sepal and

extract the petal cone with inside the anthers still joined

together (Fig.

3b, c ). Cut and open the petal cone and excise

the anthers by pushing with the tip of the forceps to the anther

ﬁ lament (Fig.

4a, b ), so avoiding to touch the anther. To place

the anthers onto the medium C, it is advisable to immerse one

tip of the forceps into the medium as the small amount of agar -

medium remaining on the tip allows to stick the anther (Fig.

4c ). The anther is placed in the 60 mm Petri dish containing

medium C (Fig.

4d ). It is extremely important to manipulate

very gently the ﬂ oral buds and, especially, the anther, avoiding

squeezing or excessive pressure, because the wounds stimulate

proliferation of somatic callus which, in turn, reduces the

androgenetic response. Detaching the whole ﬁ lament it is also

important; however, it is better to leave the ﬁ lament rather

than to risk damaging the anther, as the ﬁ lament can be easily

removed when the anther will be subcultured from medium C

to medium R.

5. Use the R medium which contains the same cytokinin of the C

medium (i.e., use R1Z for the anthers plated onto C9 medium

and R1K for C3, C6, and C13 media; see Table

1 ). Remove the

portion of the ﬁ lament which remains attached to the anther,

as well as the somatic callus developed from the anther tissue.

6. The well-formed embryos obtained in the R medium are then

transferred to V3 medium in Petri dishes as soon as they are

germinated. Plantlets are moved to a bigger container (jars or

Anther Culture in Eggplant

464

Magenta box). It is also advisable to make a backup (cuttings)

of each androgenetic plantlet by subculturing the apical shoot

with 2–3 nodes in V3 medium.

7. Gently wash out the roots from the agar and transplant the

plantlets in pots (maximum 6 cm diameter) ﬁ lled with a mix-

ture of peat (65 %), perlite (25 %), and sand (10 %). To ensure

the acclimatization of plantlets to the ex vitro conditions, the

freshly potted plantlets are maintained under high humidity

condition by covering each potted plantlet with a plastic cup

(Fig.

5c ), having 3–4 holes; the cup is then progressively

removed from the pot to reduce gradually the humidity to the

one of the growth chamber. Otherwise the freshly potted

plantlets are put in a case sealed with a transparent plastic sheet,

with holes in the top, which will be progressively removed in

10–14 days.

8. A certain percentage of plantlets will be diploid, and they can

be promptly employed in the breeding program, without the

need to double their ploidy. Diploidization is generally caused

by spontaneous chromosome doubling during the ﬁ rst micro-

spore division. Molecular analyses, using polymorphic hetero-

zygous loci markers (SSR, SNP) in the anther donor, may be

performed to conﬁ rm their gametophytic origin (i.e., having

all the loci at the homozygous state, see Fig.

7 ). It is advisable

to make this analysis if you are not completely sure that the

plantlets originated from a well-formed embryo. In fact, the

media employed for anther culture are not suitable to trigger

embryogenesis from the somatic anther tissue.

9. Use preferably young leaf for cytoﬂ uorimetric analysis.

10. Use mature and healthy leaves, cut a piece of leaf missing of

primary, secondary and tertiary veins that ensures a better

adherence of the cover slip. Observe quickly under ﬂ uorescent

microscope, after keeping the leaf in the FDA solution for a

maximum of 3–5 min. In fact, the stomata cells are the ﬁ rst

reacting to FDA, as a strong green background coloration will

hamper the chloroplast counting as soon as the FDA will pen-

etrate into the other leaf cells. In diploid and haploid plants,

the average number of chloroplasts in the stomata is about 12

and 7, respectively [

9 ].

11. Cloning of haploids by in vitro cutting increases the probabil-

ity of their conversion into the diploid status. Dissolve colchi-

cine in the lanoline by manual stirring with the aid of a spatula.

Check daily the plantlets to eliminate the young shoots com-

ing from the untreated buds, as this will further stimulate the

development of shoots from the secondary/adventitious buds

whose meristems have been exposed to the colchicine.

Giuseppe Leonardo Rotino

465

References

Fig. 7 Molecular analysis conﬁ rming the pollen origin of double- haploid plants (DHs), as they resulted homo-

zygous using a bi-allelic polymorphic SSR marker which is heterozygous in the anther donor (Hybrid); the DH

plants displayed the one or the other of the single marker present in the parental lines (A, B)

1. Research Group of Haploid Breeding (1978)

Induction of haploid plants of Solanum melon-

gena L. Proceedings of the “symposium on

plant tissue culture”, Sci. Press, Peking, p 227–

232, May 25–30

2. Isouard G, Raquin C, Demarly Y (1979)

Obtention de plantes haploides et diploides par

culture in vitro d’anthères d’aubergine ( Solanum

melongena L.). C R Acad Sci Paris 288:

987–989

3. Dumas De Vaulx R, Chambonnet D (1982)

Culture in vitro d’anthères d’aubergine

( Solanum melongena L.): stimulation de la pro-

duction de plantes au moyen de traitments à

+35°C associés à de faibles teneurs en sub-

stances de croissance. Agronomie 2:983–988

4. Chambonnet D (1985) Culture d’anthères in

vitro chez trois Solanacées maraichères: Le

piment ( Capsicum annuum L.), l’aubergine

( Solanum melongena L.), la tomate

( Lycopersicon esculentum L.) et obtention de

plantes haploides. Thèse de Docteur

d’Université, Academie de Montpellier

5. Dumas De Vaulx R, Chambonnet D, Pochard

E (1981) Culture in vitro d’anthères de piment

( Capsicum annuum L.): amélioration des taux

d’obtention de plantes chez différents géno-

types par des traitments à +35°C. Agronomie

1:859–864

6. Gu SR (1979) Plantlets from isolated pollen

culture of eggplant ( Solanum melongena L.).

Acta Bot Sin 21:30–36

7. Miyoshi K (1996) Callus induction and plant-

let formation through culture of isolated

microspores of eggplant ( Solanum melongena

L). Plant Cell Rep 15:391–395

Anther Culture in Eggplant

466

8. Corral-Martinez P, Seguì-Simarro JM (2012)

Efﬁ cient production of callus-derived doubled

haploids through isolated microspore culture

in eggplant ( Solanum melongena L.). Euphytica

187(1):47–61

9. Rotino GL (1996) Haploidy in eggplant. In:

Jain SM, Sopory SK, Veilleux RE (eds) In vitro

haploid production in higher plants. Kluwer

Academic, Dordrecht, pp 115–141

10. Salas P, Rivas-Sendra A, Prohens J, Seguí-

Simarro JM (2012) Inﬂ uence of the stage for

anther excision and heterostyly in embryogen-

esis induction from eggplant anther cultures.

Euphytica 184:235–250

11. Tuberosa R, Sanguineti MC, Conti S (1987)

Anther culture of egg-plant ( Solanum melon-

gena L.) lines and hybrids. Genet Agr 41:

267–274

Giuseppe Leonardo Rotino

467

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 26

Anther Culture in Pepper ( Capsicum annuum L.)

Verónica Parra-Vega and Jose M. Seguí-Simarro

Abstract

Anther culture is the most popular of the techniques used to induce microspore embryogenesis. This tech-

nique is well set up in a wide range of crops, including pepper. In this chapter, a protocol for anther culture

in pepper is described. The protocol presented hereby includes the steps from the selection of buds from

donor plants to the regeneration and acclimatization of doubled haploid plants derived from the embryos,

as well as a description of how to analyze the ploidy level of the regenerated plants.

Key words Androgenesis , Doubled haploid , Embryogenesis , Haploid , Microspores , Pollen , Tissue

culture

1 Introduction

Androgenesis can be deﬁ ned as the generation of an individual

derived from a nucleus of male origin, usually a haploid microspore

or young pollen grain [

1 ]. Haploid embryos or calli are produced

through the deviation of the microspore from its original gameto-

phytic pathway towards a new sporophytic pathway. Haploid

embryos may become doubled haploid individuals by themselves

or through the application of treatments for genome doubling [

2 ].

Doubled haploid individuals can be used as pure lines to produce

hybrid seeds, which reduces considerably the time and resources

needed to obtain pure lines when compared with conventional

breeding methods [

3 ].

For most of the studied species the optimal stage of male

gametophyte development to induce embryogenesis is the transi-

tion between vacuolate microspores and young bicellular pollen

[

1 , 4 ]. Technically, microspore embryogenesis can be induced

through anther culture or isolated microspore culture . Isolated

microspore culture is based on the isolation of the microspores in

liquid medium. Since the maternal tissue is removed, microspores

are directly in contact with the medium components. Therefore,

the possible formation of somatic embryo s coming from the anther

468

walls is avoided. Despite these advantages, isolated microspore

culture is more complex than anther culture and therefore it is well

set up just in a few species. Just tobacco ( Nicotiana tabacum ),

rapeseed ( Brassica napus ), wheat ( Triticum aestivum ), and barley

( Hordeum vulgare ) can be considered as model systems for micro-

spore culture [

5 ]. For most crops of agronomic interest, the most

used technique is still anther culture. Anther culture consists on

the cultivation of the anthers in a solid or semisolid medium. It can

be applied to a wide range of crops and it is the preferred technique

used to produce doubled haploid s due to its simplicity [

6 ], and to

the possibility of culturing large amounts of anthers per isolation.

In some species, the presence of the anther walls in the culture

medium seems to provide a proper environment for microspore

development, allowing for the induction of the microspores

towards embryogenesis [

7 ]. Anther culture in pepper ( Capsicum

annuum L.) has been used to produce doubled haploid plants for

breeding programs since the mid-1980s ( see Chapter

9 , this

volume).

In this chapter, a protocol for anther culture of sweet pepper is

explained according to Dumas de Vaulx et al. [

8 ] with some modi-

ﬁ cations. The protocol was adapted for commercial F1 hybrids of

sweet pepper [

9 ] and the selection of donor ﬂ ower buds was made

according to Parra-Vega et al. [

10 ]. In this protocol, the combina-

tion of two morphological markers (calix-bud length ratio and

anther pigmentation) is used to select the optimal ﬂ ower buds.

2 Materials

This stepwise protocol was developed with the following commer-

cial F1 hybrids of pepper ( C. annuum L.): ‘Herminio’ (Lamuyo

type, from Syngenta Seeds), ‘Coyote’, ‘Quito’ (California type,

both from Syngenta Seeds), and ‘Vélez’ (California type, from

Enza Zaden).

1. Plastic tubes of 50 mL.

2. Box with melting ice.

3. Laminar ﬂ ow hood.

4. Sterile Whatman paper.

5. Sterile forceps and scalpel.

6. Sterile Petri dishes 90 × 25 mm (Ø × height).

7. Paraﬁ lm.

8. Inverted or light microscope.

2.1 Plant Material

2.2 Equipment

Verónica Parra-Vega and Jose M. Seguí-Simarro

469

9. Microscope slides and cover slips.

10. Aluminum paper.

11. Incubator at 35 and 25 °C.

12. Sterile baby food jars with plastic caps.

13. Plastic plant pots 90 × 100 mm (width × height).

14. Composite soil.

15. Transparent plastic glass.

16. Growth chamber at 25 °C.

17. Pasteur pipettes, 3 mL.

18. Razor blades.

19. Filters of 30 μm pore (CellTricks, Partec).

20. Plastic tubes 3.5 mL, 55 × 12 mm (Ø × height).

21. Flow cytometer Partec Ploidy Analyzer I.

1. 70 % ethanol (v/v).

2. 4 g/L sodium hypochlorite with 0.05 % Tween (v/v).

3. Sterile distilled water (three glass jars) autoclaved at 121 °C for

20 min.

4. Induction medium: C medium (Table 1 ) supplemented with

0.01 mg/L kinetin and 0.01 mg/L 2,4-dichlorophenoxyace-

tic acid (2,4-D).

5. Regeneration medium: R medium (Table

1 ), supplemented

with 0.1 mg/L kinetin. Adjust the pH of media C and R to

5.9. Autoclave media at 121 °C for 20 min, and then pour it in

90 × 25 mm sterile Petri dishes.

6. Rooting medium: V3 medium (Table

1 ). Adjust the pH to

5.9. Autoclave medium at 121 °C for 20 min and pour it in

90 × 25 mm sterile Petri dishes and sterile baby food jars

(200 mL).

7. Lysis buffer (LB01) [

11 ]: 5 mM Tris(hydroxymethyl)amino-

methane, 2 mM Na

2 EDTA, 0.5 mM espermine, 80 mM KCl,

20 mM NaCl, 15 mM β-mercaptoethanol, and 0.1 % (v/v)

Triton X-100. The pH is adjusted at 7.5.

8. Staining buffer: 4,6-Diamidino-2-phenylindole ( DAPI )

(Partec CyStain UV precise P, PARTEC GmbH).

3 Methods

Donor plants are grown in a growth chamber at 25 °C, light inten-

sity of 200 μmol/m

2 /s with a 16 h photoperiod and 60–65 % rela-

tive humidity.

2.3 Solutions and

Culture Media

3.1 Donor Plant

Growth Conditions

Anther Culture in Pepper

470

Table 1

Macroelements, microelements, and vitamins used in the three basal

media for pepper anther culture (mg/L). C and R media from Dumas de

Vaulx et al. [

9 ]. V3 medium from Chambonnet [ 12 ]

Medium C Medium R Medium V3

Macroelements

KNO

3 2150 2150 1900

NH

4 NO

3 1238 1238 1650

MgSO

4 · 7H 2 O 412 412 370

CaCl

2 · 2H 2 O 313 313 440

KH

2 PO

4 142 142 170

Ca(NO

3 )

2 · 4H 2 O 50 50 –

NaH

2 PO

4 · H 2 O 38 38 –

(NH

4 )

2 SO

4 34 34 –

KCl 7 7 –

Microelements

MnSO

4 · H 2 O 22.130 20.130 0.076

ZnSO

4 · 7H 2 O 3.625 3.225 1.000

H

3 BO

3 3.150 1.550 1.000

KI 0.695 0.330 0.010

Na

2 MoO

4 · 2H 2 O 0.188 0.138 –

CuSO

4 · 5H 2 O 0.016 0.011 0.030

CoCl

2 · 6H 2 O 0.016 0.011 –

AlCl

3 · 6H 2 O – – 0.050

NiCl

2 · 6H 2 O – – 0.030

Vitamins and amino acids

Myo-Inositol 100.00 100.00 100.00

Pyridoxine HCl 5.500 5.500 5.500

Nicotinic acid 0.700 0.700 0.700

Thiamine HCl 0.600 0.600 0.600

Calcium pantothenate 0.500 0.500 0.500

Vitamin B12 0.030 – –

Biotin 0.005 0.005 0.005

Glycine 0.100 0.100 0.200

Chelated iron

Na

2 EDTA 18.65 18.65 37.30

FeSO

4 · 7H 2 O 13.90 13.90 27.28

Sucrose 30,000 30,000 30,000

Bacto- agar 8000 8000 8000

1. Select by eye the optimal buds for anther culture . In our geno-

types, they are covered approximately the 80 % of them by the

sepals (Fig.

1a ), according to Parra-Vega et al. [ 10 ] ( see Note

1 ). Excise the buds from the plant. Bring them to the lab in

plastic tubes immersed on melting ice ( see Note 2 ).

2. Take the buds to the laminar ﬂ ow hood.

3.2 In Vitro Culture

of Anthers

Verónica Parra-Vega and Jose M. Seguí-Simarro

471

3. Surface sterilize the buds with 70 % ethanol for 30 s, and then

with sodium hypochlorite 4 g/L for 5 min, and ﬁ nally three

washes of 4 min each with sterile distilled water ( see Note 3 ).

4. Place the buds over sterile Whatman paper and excise them to

extract the anthers ( see Note 4 ). At this step, make a second

selection of the buds. Culture only buds containing anthers

with purple distal tips (Fig.

1e ), according to Parra-Vega et al.

[

10 ]. In case the optimal stage of anther development has not

been well set up in advance for the genotype used, it is highly

recommended, at this point, to check the microspores/ pollen

stage of every bud before culturing them ( see Note 5 ).

5. Place the selected anthers in Petri dishes with C medium. Place

them with their concave part in contact with the medium. Seal

the dishes with Paraﬁ lm and introduce them in the incubator

at 35 °C in darkness for 4 days ( see Note 6 ).

6. At day 4, place the dishes in the incubator at 25 °C with a 12-h

photoperiod for 4 days more.

7. At day 8, transfer the anthers to R medium and incubate them

at 25 °C, light intensity of 32 μmol/m

2 /s and a 12 h photope-

riod. Every 2 months, change the anthers to fresh R medium.

8. As soon as the embryos pop out of the anthers, pick them with

forceps and transfer them to V3 medium in 90 × 25 mm Petri

Fig. 1 Process of anther culture in pepper . ( a ) Flower bud at the right stage for anther isolation. ( b – d ) Anther

extraction out of the bud: transversal cut of the ﬂ ower bud ( b ), longitudinal cut of the ﬂ ower bud surface ( c ),

and opening of the ﬂ ower bud with scalpel and forceps to extract the anthers ( d ). ( e ) Anthers at the right stage

for isolation. White arrow points the right position to culture the anthers in medium ( concave part ). ( f ) Anther

culture d in vitro producing two microspore -derived embryos ( white arrows ) in c medium. ( g ) Microspore -

derived embryo germinated in V3 medium. ( h ) Microspore-derived seedling cultured in vitro in V3 medium. ( i )

Acclimated seedling cultured ex vitro in a plastic plant pot. Bars: a – e , 2 mm; f and g , 5 mm; h , 1 cm; i , 2 c m

Anther Culture in Pepper

472

dishes, incubate them at 25 °C, light intensity of 32 μmol/

m

2 /s and a 12 h photoperiod. Transfer the embryos that ger-

minate correctly to sterile baby food jars with V3 medium ( see

Note 7 ).

9. When seedlings develop a proper root system (one or two pri-

mary roots and some secondary roots), transfer them to plastic

plant pots with wet soil.

10. Acclimate the seedlings in the growth chamber at 25 °C and a

16 h photoperiod ( see Note 8 ).

1. Analyze the nuclear DNA content with a ﬂ ow cytometer

(Partec Ploidy Analyzer I) according to its commercial speciﬁ -

cations. Use DAPI as the ﬂ uorescent stain.

2. Use donor plants as control for 2C DNA content. Plants

derived from embryos will be analyzed in order to know the

ploidy level ( see Note 9 ).

3. Excise young leafs from the plant and place them in a box with

ice ( see Note 10 ).

4. Chop with a razor blade a piece of 1 cm

2 of a young leaf in a

plastic Petri dish containing 0.5 mL of lysis buffer ( see Note

11 ).

5. Filter the extracted nuclei with a 30 μm pore ﬁ lter into a

3.5 mL plastic tube.

6. Add 1.5 mL of DAPI staining buffer with a 3 mL Pasteur

pipette.

7. Keep the tubes on ice for 2 min prior to analyze the samples

using the ﬂ ow cytometer. Count a minimum of 10,000 cells

per sample.

4 Notes

1. The selection of anthers is one of the critical steps of anther

culture . The anthers must contain vacuolate microspores and

young bicellular pollen grains to efﬁ ciently induce embryogen-

esis. As this parameter determination is highly genotype depen-

dent, it is recommended to study previously, in each genotype,

the right size and appearance of anthers containing the appro-

priate stage of microspore /pollen development to be induced

towards embryogenesis.

2. Once the buds are excised from the plant, keep them on ice in

order to slow down the development of the microspores/ pollen .

Also, keep the sterilized solutions at 4 °C before using them to

reduce the degradation process of anthers.

3.3 Analysis of

the Ploidy Level

Verónica Parra-Vega and Jose M. Seguí-Simarro

473

3. Pour the sterilized solutions into the plastic tube, close the lid

and shake the solutions during the corresponding time for

each solution. After that, open the lid and remove the liquid

keeping the buds. Pour the next solution into the tube and

repeat the process. An alternative to the plastic tubes is to use

tea ﬁ lter sieves.

4. Excise the anthers with a scalpel avoiding breaking them. First,

make a transversal cut at the basal part of the bud (near to the

pedicel), removing the basal part of the ﬂ oral bud (Fig.

1b ).

Second, make a longitudinal cut, only at the surface of the bud

(Fig.

1c ), to open the sepals. Later, take away the sepals and

petals with forceps, and extract the anthers (Fig.

1d ). It is

important to remove the anther ﬁ lament as much as possible,

just to avoid callus formation from this tissue, which is espe-

cially prone to proliferate.

5. After extracting the anthers from the bud, take one anther to

observe it under the microscope and keep the remaining

anthers waiting in the laminar ﬂ ow hood. Place the anther

onto a microscope slide with a drop of water, chop the anther

with a razor blade in order to extract the microspores/ pollen

and cover it with a standard cover slip. Observe the prepara-

tion under a light or inverted microscope checking the stages

of microspores contained. If the anther contains mostly vacu-

olate microspores and young bicellular pollen, the rest of

anthers from the same bud will be used for anther culture .

6. Cover the Petri dishes with aluminum paper to create a dark-

ness environment inside the incubator.

7. Transfer the germinated embryos to baby jars in order to

increase the space to develop the roots and aerial parts of the

new plant.

8. In order to avoid drastic change in humidity conditions, use a

transparent plastic cup to protect the seedlings. Pinch holes in

the cup every 2 days, to gradually reduce the humidity inside

the cup down to the levels of the growth chamber. Then

remove the glass.

9. The ﬂ ow cytometer is used to analyze the ploidy level, but

when a 2C individual appears, molecular analysis marker (pref-

erentially SSRs) has to be performed in order to clarify whether

this individual has a somatic or an embryogenic origin. For

donor plants polymorphic for the SSR used, if the regenerated

samples analyzed are homozygous for the used molecular

markers, the origin of these plants will be gametophytic.

However, if the samples are heterozygous for the SSRs used,

their origin will likely be somatic (most likely coming from

anther wall tissues).

Anther Culture in Pepper

474

10. Young tissues are used to analyze the ploidy level because these

tissues present more cells in G2 phase; therefore the second

peak of the histogram appears clearer.

11. The nucleic extraction buffer from Partec (CyStain UV precise

P, PARTEC GmbH) may be used at this step. However, with

pepper is recommended to use the lysis buffer in order to slow

down the oxidizing process of pepper samples.

Acknowledgments

This work was supported by the grant AGL2014-55177 from

Spanish MINECO to J.M.S.S.

References

1. Seguí-Simarro JM (2010) Androgenesis revis-

ited. Bot Rev 76:377–404

2. Seguí-Simarro JM, Nuez F (2008) Pathways to

doubled haploidy: chromosome doubling dur-

ing androgenesis. Cytogenet Genome Res

120:358–369

3. Wedzony M, Forster BP, Zur I, Golemiec E,

Szechynska-Hebda M, Dubas E, Gotebiowska

G (2009) Progress in doubled haploid technol-

ogy in higher plants. In: Touraev A, Forster BP,

Jain SM (eds) Advances in haploid production

in higher plants. Springer, Dordrecht,

Netherlands, pp 1–33

4. Soriano M, Li H, Boutilier K (2013)

Microspore embryogenesis: establishment of

embryo identity and pattern in culture. Plant

Reprod 26:181–196

5. Forster BP, Heberle-Bors E, Kasha KJ, Touraev

A (2007) The resurgence of haploids in higher

plants. Trends Plant Sci 12:368–375

6. Germanà MA (2011) Anther culture for hap-

loid and doubled haploid production. Plant

Cell Tissue Organ Cult 104:283–300

7. Seguí-Simarro JM, Nuez F (2008) How micro-

spores transform into haploid embryos:

changes associated with embryogenesis

induction and microspore-derived embryogen-

esis. Physiol Plant 134:1–12

8. Dumas de Vaulx R, Chambonnet D, Pochard E

(1981) Culture in vitro d’anthères de piment

( Capsicum annuum L.): amèlioration des taux

d'obtenction de plantes chez différents géno-

types par des traitments à +35°C. Agronomie

1:859–864

9. Parra-Vega V, Renau-Morata B, Sifres A, Seguí-

Simarro JM (2013) Stress treatments and

in vitro culture conditions inﬂ uence micro-

spore embryogenesis and growth of callus from

anther walls of sweet pepper ( Capsicum ann-

uum L.). Plant Cell Tissue Organ Cult

112:353–360

10. Parra-Vega V, González-García B, Seguí-

Simarro JM (2013) Morphological markers to

correlate bud and anther development with

microsporogenesis and microgametogenesis in

pepper ( Capsicum annuum L. ). Acta Physiol

Plant 35:627–633

11. Dolezel J, Binarova P, Lucretti S (1989)

Analysis of nuclear DNA content in plant

cells by flow cytometry. Biol Plant

31:113–120

12. Chambonnet D (1988) Production of hap-

loid eggplant plants. Bulletin interne de la

Station d'Amelioration des Plantes

Maraicheres d'Avignon-Montfavet, France,

1-10

Verónica Parra-Vega and Jose M. Seguí-Simarro

475

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 27

Microspore Embryogenesis Through Anther Culture

in Citrus clementina Hort. ex Tan.

Benedetta Chiancone and Maria Antonietta Germanà

Abstract

Anther culture is a biotechnological method that allows to obtain, in one step, homozygous plants, very

important to plant breeding, due to their numerous applications in mutation research, selection, genome

sequencing, genetic analysis, and transformation. To induce the microspores, i.e., the immature male gam-

etes, to switch from the normal gametophytic pathway to the sporophytic one, it is necessary to submit

them to a type of stress, such as high or low temperature, starvation, or magnetic ﬁ eld. Stress can be

applied to the donor plants and/or the ﬂ oral buds or the anthers or the isolated microspores, before or

during the culture. In this chapter, the protocol to induce gametic embryogenesis from anther culture of

several cultivars of Citrus clementina Hort. ex Tan. is reported.

Key words Anther culture , Citrus , Clementine , Doubled haploid s , Gametic embryogenesis , Isolated

microspore culture , Microspore embryogenesis , Somatic embryo genesis

1 Introduction

The conventional methods applied to Citrus breeding are time-

consuming and limited by many factors. Biotechnological meth-

ods, and, among them, haploidy technology, are a valuable support

to increase the efﬁ ciency and to speed up the breeding programs.

The interest of breeders in haploids and doubled haploid s relies

mainly in the possibility of obtaining homozygosity in one step,

particularly in woody plants, generally characterized by a long

reproductive cycle, a high degree of heterozygosity, large size, and,

sometimes, by self-incompatibility [

1 – 3 ]. Haploid technology is

important for its potential use in mutation research, selection,

genetic analysis, transformation, and the production of homozy-

gous cultivars. Moreover, in Citrus , where somatic hybrid ization is

a well-established protocol, haploid protoplasts can be fused with

diploid ones in order to obtain triploids, which are particularly

important since they are seedless [

2 – 4 ].

476

In Citrus , the ﬁ rst haploid seedlings were obtained by the

application of gamma rays in Citrus natsudaidai [

5 ]. After that,

many studies have been carried out on gametic embryogenesis to

obtain haploid and doubled haploid plants, through anther cul-

ture , but not much of them have been successful. For example,

only heterozygous plantlets have been obtained by anther culture

in C. aurantium [

6 , 7 ], C. aurantifolia [ 8 ], C. madurensis [ 9 ],

C. reticulata [

10 ], Poncirus trifoliata [ 11 , 12 ], and C. sinensis [ 13 ,

14 ]. Haploid plantlets have been recovered from Poncirus trifoli-

ata [

15 ] and C. madurensis [ 16 ]; one doubled haploid plantlet has

been obtained from the hybrid No. 14 of C. ichangensis × C. retic-

ulata [

12 ]; haploid but albino embryoids of Mapo tangelo C. deli-

ciosa × C. paradisi [

17 ], haploid and diploid calli, embryoids and

leafy structures but no green plants of C. limon [

18 ], and haploid

embryoids of Clausena excavata [

19 ] have been also achieved.

Furthermore, haploid, doubled haploid and triploid plantlets, and

highly embryogenic calli of C. clementina Hort. ex Tan. were

recovered [

10 , 14 , 20 – 23 ].

To induce gametic embryogenesis , it is necessary to switch

microspore development from the gametophytic to the sporo-

phytic pathway, usually subjecting microspores to a stress treat-

ment [

2 , 6 , 24 , 25 , 26 ]. Stress can be provided through the

growing conditions of the donor plants and/or as treatments

applied to the ﬂ oral buds or to the anther or to the isolated micro-

spores, before or during the culture. Actually, all aspects of the in

vitro culture protocol could be classiﬁ ed as stresses [

27 ]. The stress

seems to act by altering the polarity of the division at the ﬁ rst hap-

loid mitosis, involving reorganization of the cytoskeleton [

28 ],

delaying and modifying pollen mitosis, blocking starch produc-

tion, or dissolving microtubules [

29 ].

Also in Citrus , numerous studies were conducted to obtain

regeneration through anther and isolated microspore culture

techniques, testing the microspore response to different stress

treatments applied before and after the culture [

4 , 10 , 17 , 18 , 23 ,

24 , 30 , 31 ]. In particular, in Citrus clementina Hort. ex Tan.,

several stress treatments have been tested to induce microspore

embryogenesis , both by isolated microspore and by anther cul-

ture , particularly low- and high-temperature pretreatments and

magnetic ﬁ eld treatments (

10 , 13 , 14 , 20 – 23 , 32 , 33 , and unpub-

lished results).

In this chapter, the protocol successfully used to induce micro-

spore embryogenesis through anther culture in several Citrus cle-

mentina Hort. ex Tan. cultivars is reported [

13 , 14 , 22 , 23 ].

Benedetta Chiancone and Maria Antonietta Germanà

477

2 Anther Culture

Immature ﬂ ower buds of Citrus clementina Hort. ex Tan., cvs.

Nules, SRA 63, Monreal, Corsica, and Hernandina, with anthers

containing microspores at the vacuolated stage of development,

collected from ﬁ eld growing trees.

1. Stereo microscope, light microscope, ﬂ uorescent microscope,

slides.

2. Laminar ﬂ ow hood, forceps, scalpels, glass bead sterilizer or

burners.

3. Plastic/glass 1000 mL beakers, 1000 mL graduated cylinders,

Petri dishes (60 mm diameter tissue culture Petri dishes),

Magenta boxes (Sigma V8505), test tubes, 1000 mL screw-

capped Pyrex bottles, Paraﬁ lm, magnetic stirrers, spin bars,

100 and 1000 μL micropipettes, and micropipette tips.

4. pH meter.

5. Autoclave.

6. Jiffypots, peat moss, sand, soil, polythene bags.

1. Sterile distilled water, 70 % (v/v) ethyl alcohol, 20 % (v/v)

commercial bleach.

2. 1 mg/mL of 4′,6-diamidino-2-phenylindole dihydrochloride

( DAPI ).

3. In Table

1 , media used for inducing gametic embryogenesis

(IM), for embryogenic calli proliferation (PM), and for embryo

germination (GM) are reported ( see Note 1 ).

1. Collect unopened ﬂ owers 3.5–5.0 mm long, from plants in

February to April, depending on the season and on the geno-

type (Fig.

1a ).

2. Store ﬂ ower buds at the immature stage at 4 °C for about

1 week ( see Note 2 ).

1. Determine the pollen development stage, staining one or more

anthers per ﬂ oral bud size with DAPI .

2. Squash anthers in few drops of DAPI solution (1 mg/mL) and

observe slides under a ﬂ uorescent microscope to identify the

pollen development stage ( see Note 3 ).

1. To prepare a ﬁ nale volume of 1 L, start with 500 mL of dis-

tilled water containing a magnetic stirrer ( see Note 4 ).

2. Start adding salts and vitamin mixture, then the carbon source

and the growth regulators, mixing properly.

2.1 Materials

2.1.1 Plant Material

2.1.2 Equipment

2.1.3 Solutions

and Culture Media

2.2 Methods

2.2.1 Flower Bud

Collection

2.2.2 DAPI Staining

and Developmental Stage

Determination

2.2.3 Culture Medium

Preparation

and Sterilization

Anther Culture in Citrus clementina

478

Table 1

Media composition used for Citrus clementina Hort. ex Tan. anther culture

Components

IM PM GM

Per liter

N6 Chu basal salts 1×

MS basal salts – 1× 1×

N&N vitamins 1×

MS vitamins 1× 1×

Galactose 18 g

Lactose 36 g

Sucrose 50 g 30 g

Ascorbic acid 500 mg 500 mg 500 mg

Myo-Inositol 5 g

Biotin 500 mg

Thiamine 5 mg

Pyridoxine 5 mg

Casein 500 mg

Glycine 2 mg

Glutamine 800 mg

Serine 100 mg

Malt extract 500 mg 500 mg 500 mg

Coconut water 100 mL

6-Benzylaminopurine 0.5 mg

2,4-Dichlorophenoxyacetic acid 0.5 mg

Gibberellic acid 0.5 mg 1 mg

Kinetin 0.5 mg

1-Naphthaleneacetic acid – 0.02 mg 0.01 mg

Thidiazuron 0.5 mg

Zeatin 0.5 mg

Agar 8.5 g 8.0 g 7.5 g

pH 5.8 5.8 5.8

Abbreviations : MS = [ 41 ]; N&N = [ 42 ]; N6 = [ 43 ]; IM = induction medium; PM = prolif-

eration medium; GM = germination medium

Benedetta Chiancone and Maria Antonietta Germanà

479

3. Adjust the pH of media to 5.8, with 1 N KOH or 1 N HCl,

and then bring to volume adding distilled water till 1 L.

4. Add agar directly in the bottle before the medium, without

mixing ( see Note 5 ).

5. Put the bottle, without closing completely the cap, in the auto-

clave and sterilize it at 110 kPa, 121 °C for 20 min.

6. Under the laminar ﬂ ow hood, pour in 60 mm Petri dishes the

medium, only when its temperature is lower than 60 °C.

1. Under the laminar ﬂ ow hood, to sterilize ﬂ ower buds, immerse

them, ﬁ rstly for 3 min in 70 % (v/v) ethyl alcohol and then in

25 % commercial bleach solution (about 1.5 % active chlorine

in water) with few drops of Tween 20, for 15–20 min. Finally,

rinse them three times with sterile distilled water.

2. Isolate anthers, ﬁ rst by removing the petals and then separat-

ing them from stamens (Fig.

1b ).

3. Put 60–80 anthers per each Petri dish containing 10 mL of

induction medium (IM) (Table

1 ).

4. Use Paraﬁ lm to seal Petri dishes, before the incubation at

27 ± 1 °C, in the dark, for the ﬁ rst month and then under cool

white ﬂ uorescent lamps (Philips TLM 30W/84) with a photo-

synthetic photon ﬂ ux density of 35 μmol m

-2 s

-1 and a 16 h

light photoperiod.

5. Observe the cultures for 10 months, every 2 weeks.

1. Once embryos and embryogenic calli start to appear (after 2–3

months), transfer them to proliferation medium (PM) ( see

Note 6 ) (Fig.

2a, b ).

2. Subculture the stock culture lines every 45 days, keeping them

at the same light and temperature conditions.

2.2.4 Flower Bud

Sterilization, Anther

Isolation, and Culture

2.2.5 Embryogenic

Callus Maintenance

Fig. 1 Plant material. ( a ) Citrus clementina Hort. ex Tan., cv SRA 63 ﬂ ower bud (5 mm) at the suitable develop-

mental stage. ( b ) Anther of Citrus clementina Hort. ex Tan., cv Nules containing microspores at the vacuolate

developmental stage ( c ) Bar = 10 μm

Anther Culture in Citrus clementina

480

1. Isolate the well- developed embryos and culture in 100 mm

Petri dishes containing the germination medium (GM) (Fig.

2c ) (Table 1 ) ( see Note 7 ).

2. Keep the culture in the light at 27 ± 1 °C (with a 16 h

photoperiod).

3. Move germinated embryos in Magenta boxes or in test tubes

containing the same medium, with 5–6 week subcultures

( see Note 8 ) (Fig.

2d ).

1. Wash the rooting apparatus of well-developed plantlets with

sterile distilled water to remove the medium residues.

2. Transplant plantlets, 4–5 cm high, in Jiffypots or in pots con-

taining sterile peat moss, sand, and soil, in the ratio of 1:1:1,

and grow them in the greenhouse.

3. To avoid dehydration, cover the plantlets with polythene bags

for the ﬁ rst 40–50 days ( see Note 9 ).

2.2.6 Embryo

Germination

2.2.7 Plant Development

and Acclimatization

Fig. 2 Microspore embryogenesis through anther culture . ( a ) Direct embryogenesis from an anther of Citrus

clementina Hort. ex Tan., cv Monreal. ( b ) Embryogenic callus production from an anther of Citrus clementina

Hort. ex Tan., cv Nules, after 3 months of culture. ( c ) Microspore-derived embryos of Citrus clementina Hort. ex

Tan., cv Hernandina. ( d ) Plantlet of Citrus clementina Hort. ex Tan., cv Corsica, regenerated from anther culture

and transferred to test tube

Benedetta Chiancone and Maria Antonietta Germanà

481

3 Regenerant Characterization

1. A portion of 0.5 cm

2 leaf tissue collected from a regenerated

plantlet (or the equivalent part of a regenerated embryo) and

the same amount of a mother plant young leaf.

2. Razor blade, nylon gauze ﬁ lter (Partec CellTrics

® ).

3. Extraction buffer (Partec CyStain

® UV Precise); staining buffer

(Partec CyStain

® UV Precise).

4. Flow cytometer (Partec, Münster, Germany).

1. Cut by a razor blade a portion of 0.5 cm

2 leaf tissue collected

from a regenerated plantlet (or the equivalent part of a regen-

erated embryo) and the same amount of a mother plant young

leaf. Chop them together in 1 mL of extraction buffer (Partec

CyStain

® UV Precise), to release the nuclei from the cells.

2. Use 30 μm nylon gauze ﬁ lter (Partec CellTrics

® ) to remove debris.

3. Add the staining buffer (Partec CyStain

® UV Precise) to the

suspension.

4. Inject the suspension in the ﬂ ow cytometer (Partec, Münster,

Germany) to determine relative DNA content of the samples

(Fig.

3a ) ( see Note 10 ).

3.1 Ploidy Analysis

of Regenerants

by Flow Cytometer:

Materials and Method

3.1.1 Materials

3.1.2 Methods

Peak Index Mean Area Area% CV% ChiSqu

1 1.000 100.15 1186 36.46 4.28 1.11

2 1.507

400

320

240

Counts

160

80

0

0 100 200 300

FL4

400 500

3n

2n

150.93 2058 63.54 3.72 1.11

ab

Fig. 3 Characterization of anther culture regenerants. ( a ) Cytoﬂ uorimetric analysis: histograms of ﬂ uorescence

intensity of a diploid control ( C. clementina Hort. ex Tan. mother plant) and of a triploid regenerant of C. clem-

entina Hort. ex Tan., cv Corsica, obtained by anther culture. ( b ) Microsatellite analysis: polyacrylamide gel

electrophoresis of microsatellites TAA15 showing the homozygosity of a regenerant from anther culture of C.

clementina Hort. ex Tan., cv Nules. DNA was extracted from leaves of mother plants (P) and of one anther

culture regenerant (R)

Anther Culture in Citrus clementina

482

Microsatellite analysis has several applications in Citrus breeding.

For example, in the case of anther culture , because it is possible to

regenerate from the somatic anther tissue, as well as from the

microspores, microsatellites , being codominant markers, allow to

discriminate heterozygous and homozygous regenerants (Fig.

3b )

( see Note 11 ).

DNA extraction

1. Sterile Eppendorf tubes, sterile 100 and 1000 μL tips, 100 and

1000 μL micropipettes, centrifuge, vortex, timer, liquid nitro-

gen container, gloves, sterile pestles, biosafety cabinet, water

bath.

2. Ethyl alcohol, liquid nitrogen , phenol, ammonium acetate,

isopropanol.

3. Extraction buffer (EB) (stocks 100 mM Tris–HCl pH 8, 50 mM

Na

2 EDTA pH 8, 500 mM NaCl, 10 mM β-mercaptoethanol,

3 % sodium dodecyl sulfate, SDS).

4. TE buffer [10 mL of 1 M Tris–HCl (pH 8.0), 2 mL EDTA

(0.5 M), Milli-Q water to 1000 mL].

DNA ampliﬁ cation

1. PT 100 thermal cycler (MJ Research, USA).

2. 0.5 mL Eppendorf tubes, micropipettes (1–20 and 20–200

μL), gloves, crushed ice.

3. Primers (such as TAA1, TAA15, TAA41, TAA 45, [

34 ]),

dNTPs, template DNA, Taq DNA polymerase.

4. 10× PCR buffer (500 mM KCl, 15 mM mgCl

2 , 100 mM Tris–

HCl, pH 8.3).

Polyacrylamide ( PA ) gel electrophoresis

1. Gloves and polyacrylamide gel electrophoresis system.

2. PA mixture: 6 % acrylamide solution, 50 μL N , N , N ′, N ′-

tetramethylethylenediamine (TEMED), 600 μL 10 % ammo-

nium persulphate (APS).

Silver staining

1. Polyacrylamide gel, tray.

2. Fixer (10 % acetic acid): 50 mL glacial acetic acid in 450 mL

distilled water.

3. Silver stain: 3 mL 1 N silver nitrate solution; 500 mL distilled

water; sodium thiosulphate solution (0.1 N), formamide.

4. Developer: 15 g sodium carbonate; 500 mL distilled water and

put it at 4 °C, 75 mL of sodium thiosulphate solution (0.1 N),

and 0.75 mL of formamide.

3.2 Molecular

Characterization:

Microsatellite Analysis

3.2.1 Materials

Benedetta Chiancone and Maria Antonietta Germanà

483

DNA extraction

1. Isolate a young leaﬂ et or 150 mg of callus from each regenerant

and from the mother plant; process each sample separately.

2. Warm up the EB on the 37 °C water bath, under sterile bio-

safety cabinet.

3. Cool the centrifuge to 4 °C.

4. Put the sample in the Eppendorf tube and reduce it in powder,

adding liquid nitrogen .

5. Add 700 μL of EB, vortex, incubate at 65 °C for 10 min, and

centrifuge for 5 min at 13,000 rpm (16,060 RCF).

6. Transfer the supernatant in a clean Eppendorf tube, add 700

μL of phenol, and vortex for few seconds.

7. After centrifuging for 5 min at 13,000 rpm, transfer the super-

natant in a clean Eppendorf tube.

8. Add 65 μL of ammonium acetate (NH

4

+ Ac) and 450 μL of

cold isopropanol and mix lightly.

9. Centrifuge for 10 min at 13,000 rpm, eliminate the superna-

tant, and add 700 μL of 70–75 % of cold ethyl alcohol.

10. Centrifuge for 10 min at 13,000 rpm and eliminate the ethyl

alcohol, leaving uncovered the Eppendorf tubes.

11. Resuspend DNA in 100 μL of TE buffer and store at 4 °C for

one night.

12. Quantify or store at −20 °C.

DNA ampliﬁ cation

1. In a 0.5 mL Eppendorf tube on ice, add all the reagents in the

following order: 30 μL sterile distilled H

2 O, 5 μL 10× PCR

buffer, 4 μL dNTP Mix (1.25 mM), 2.5 μL per each primer, 4

μL MgCl

2 (25 mM), and just before the reaction starts, add

the Taq DNA polymerase (Amersham Biosciences, USA) ( see

Note 12 ).

2. Add 15 μL of cocktail to the genomic DNA.

3. Place the Eppendorf tubes in the thermocycler. Use the PCR

thermal proﬁ le: 94 °C for 5 min for 1 cycle; 94 °C for 60 s, 55

°C for 30 s, 72 °C for 60 s for 32 cycles, 72 °C for 5 min [

21 ].

Polyacrylamide ( PA ) gel electrophoresis

1. Place 0.4 mm spacers on glass plates and pour the acrylamide

mixture between the plates using a syringe, until solution ﬁ lls

the space between the plates ( see Note 13 ), and then lay the

plates ﬂ at.

2. Insert comb teeth up and clamp, and then leave it to polymer-

ize for 30–45 min.

3.2.2 Methods

Anther Culture in Citrus clementina

484

3. Put the plates in the apparatus. Add in the chambers warm

0.5× TBE (±1 cm over the shorter glass), and pre-run the gel

at low wattage for 10 min (40 °C).

4. Load the samples and connect the apparatus: 40 °C, 40 W

constant ± 2 h.

5. Stop the running, open the circuit, and eliminate TBE from

upper chamber.

6. Open the glass plates, remove the spacers, and stain gel with

silver staining.

Silver staining

1. Leave the gel in the ﬁ xer for 30 min, and then wash the gel two

times, 10–15 min each time.

2. Immerse the gel in the silver stain for 30 min.

3. Just before the gel developing, add the cold developer

solution.

4. Agitate the silver stain for 10 s, before eliminating it, and then

add the developer.

5. Wait the band development, and then add the ﬁ xer.

6. Wash the gel with water for 20 min, and then put it vertically

to dry.

7. Photograph or scan the gel for observations.

4 Notes

1. Culture medium composition is one of the crucial factors

affecting the gametic embryogenesis induction. In the last 20

years, several culture media have been used in Citrus clemen-

tina Hort. ex Tan. in vitro anther culture . In particular, experi-

ments were carried out testing the effect of the addition of

different carbon sources [

10 ] and gelifying agents (data not

published) or of different growth regulator combinations,

including thidiazuron [

21 ] and polyamines [ 36 ].

2. Storage at 4 °C has the double function of preserving the

ﬂ ower buds from senescence and of stressing the microspores

before the culture.

3. Several studies report that in Citrus clementina Hort. ex Tan.,

ﬂ ower buds of 3.5–5.0 mm size contain the highest ratio of

uninucleated/vacuolated microspores (Fig.

1c ) [ 1 – 4 ].

4. Starting with a lower volume guarantees to not overcome the

ﬁ nal 1 L volume.

5. Agar is not dissolvable at room temperature.

Benedetta Chiancone and Maria Antonietta Germanà

485

6. Anthers require 2–3 months to initiate callus and embryo pro-

duction. Many investigations report that most of the calli

obtained in Citrus anther culture are non-morphogenic, but

some of them appear friable and white and differentiate into a

clump of embryos. This type of callus is highly embryogenic,

and its embryogenic potential is maintained for several years.

From only one anther, it is possible to obtain a high amount of

embryogenic callus and more than 100 embryos after several

subcultures [

22 ].

7. In the germination medium (GM), the microspore -derived

embryos follow the same developmental steps of the zygotic

ones: globular, heart, torpedo, and cotyledonary stages.

Furthermore, secondary embryogenesis can be observed, more

frequently in the root region of the embryos. Sometimes, tera-

tomatal structures and morphological anomalies, cotyledonary-

fused, pluricotyledonary, or fascinated and thickened embryos,

are observed [

35 ].

8. It is expected that 80–89 % of the cultured embryos will ger-

minate in vitro [

36 ].

9. In order to reduce the humidity level, it is recommended to

make some holes in the plastic bag and gradually increase their

sizes.

10. In Citrus clementina Hort. ex Tan., through anther culture ,

regeneration of calli and plantlets of different ploidy levels,

haploid , doubled haploid , triploid, tetraploid, aneuploid,

and mixoploid, have been reported, with a preponderance of

triploids (around 80 %) [

22 ]. The obtaining of non-haploids

may be due to the regeneration from the anther walls to the

fusion of several nuclei, to the endomitosis within the pollen

grain, and to meiotic irregularities in the microspores

[

37 – 40 ].

11. In Citrus , several microsatellites , such as TAA 1, TAA 15,

TAA27, TAA 33, TAA 41, TAA 45, TAA 52, CAGG 9, and

CAC23, were used [

34 ]. It is needed to individuate which

microsatellites are heterozygous in the mother plant. The pres-

ence of one band in the regenerant and two bands in the

mother plant is considered the conﬁ rmation of the gametic

origin of the regenerant.

12. To have enough solution, the dose of each component has to

be multiplied for the number of samples plus 1; Taq DNA

polymerase has to be kept on ice.

13. It is important to avoid the bubble formation in the gel; for

this reason, it is worth to invert the syringe to expel any

trapped air.

Anther Culture in Citrus clementina

486

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34. Kijas JMH, Thomas MR, Fowler JCS, Roose

ML (1997) Integration of trinucleotide micro-

satellites into a linkage map of Citrus . Theor

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35. Benelli C, Germanà MA, Ganino T, Beghè D,

Fabbri A (2010) Morphological and anatomi-

cal observations of abnormal somatic embryos

from anther cultures of Citrus reticulata . Biol

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36. Chiancone B, Germanà MA (2013)

Micropropagation of Citrus spp. by

Organogenesis and Somatic Embryogenesis.

In: Lambardi M, Ozudogru EA, Jain SM (eds)

Protocols for micropropagation of selected

economically-important horticultural plants.

Humana Press-Springer, Berlin, pp 99–118

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Anther Culture in Citrus clementina

Part V

Stepwise Protocols on Pivotal Topics

491

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 28

Detection of Epigenetic Modiﬁ cations During Microspore

Embryogenesis: Analysis of DNA Methylation Patterns

Dynamics

Pilar S. Testillano and María Carmen Risueño

Abstract

Methylation of 5-deoxy-cytidines of DNA constitutes a prominent epigenetic modiﬁ cation of the chromatin

ﬁ ber which is locked in a transcriptionally inactive conformation. Changes in global DNA methylation are

involved in many plant developmental processes during proliferation and differentiation events. The analy-

sis of the changes of global DNA methylation distribution patterns during microspore embryogenesis

induction and progression will inform on the regulatory mechanisms of the process, helping in the design

of protocols to improve its efﬁ ciency in different species. To investigate the DNA methylation dynamics

during microspore embryogenesis in the different cell types present in the cultures, the analysis of spatial

and temporal pattern of nuclear distribution of 5-methyl-deoxy-cytidine (5mdC) constitutes a potent

approach. The immunolocalization of 5mdC on sections and subsequent confocal laser microscopy

analysis have been developed for in situ cellular analysis of a variety of plant samples, including embryo-

genic microspore and anther cultures. Quantiﬁ cation of 5mdC immunoﬂ uorescence intensity by image

analysis software also permits to estimate differences in global DNA methylation levels among different cell

types during development.

Key words Anther culture , Confocal laser scanning microscopy , Embryo , Epigenetics ,

Immunoﬂ uorescence , 5-Methyl-deoxy-cytidine , Microspore culture , Pollen

1 Introduction

Plant developmental processes, as differentiation and proliferation,

are accompanied by chromatin remodeling and epigenetic repro-

gramming. DNA methylation constitutes a prominent epigenetic

modiﬁ cation of the chromatin ﬁ ber which is locked in a transcrip-

tionally inactive conformation [

1 ]. Microspore embryogenesis can

be induced either in anther or isolated microspore culture s [

2 , 3 ];

in these in vitro systems, after a stress treatment, microspores are

reprogrammed and change their gametophytic developmental

pathway toward embryogenesis. Nevertheless, this process presents

a low efﬁ ciency in many species because its regulatory mechanisms

492

are not well known. Stress-induced plant cell reprogramming

involves changes in global genome organization, being the epigen-

etic modiﬁ cations, DNA methylation and histone modiﬁ cations ,

key factors in the regulation of genome ﬂ exibility. Therefore, the

analysis of the epigenetics modiﬁ cations involved in microspore

embryogenesis induction will inform on its regulatory mechanisms

and open the door to exploit the process more efﬁ ciently for plant

breeding and biotechnology purposes in agriculture, selection and

propagation of forest resources, and environment control.

Research in the past years has revealed exciting ﬁ ndings with

regard to epigenetic mechanisms controlling plant developmental

processes [

4 ]. However, the knowledge of the DNA methylation

and histone modiﬁ cation regulation during relevant developmen-

tal programs in ﬂ owering plants, such as gametogenesis or embryo-

genesis, is limited [

4 – 8 ]. Difﬁ cultly in accessing speciﬁ c cell types

inside the very young embryo or endosperm inside the maternal

tissues or the developing microspores inside the anthers has made

biochemical and molecular analysis sometime problematic.

Although partially overcome by the use of in vitro systems, in situ

localization approaches using modern bioimaging technology have

become essential tools [

9 – 11 ]. To investigate the global DNA

methylation dynamics during plant embryogenesis, the analysis of

spatial and temporal pattern of nuclear distribution of 5-methyl-

deoxy-cytidine (5mdC) constitutes a potent approach, which per-

mitted to distinguish among cell types in the same embryo, in

comparison with the electrophoretic and ELISA assays used to

quantify the percentage of methylated cytidines in genomic DNA.

Immunolocalization of 5mdC and confocal analysis have been

developed to several plant cell types, tissues, and organs [

5 – 7 , 12 ,

13 ], and the results demonstrate the versatility and feasibility of the

approach for different plant samples, revealing deﬁ ned DNA meth-

ylation nuclear patterns associated with differentiation and prolif-

eration events of various plant cell types and developmental

programs. Quantiﬁ cation of 5mdC immunoﬂ uorescence intensity

by appropriate confocal image software also permits to estimate

differences in global DNA methylation levels among different cell

types of the same organ during development and under different

physiological conditions. During microspore embryogenesis , the

analysis of the dynamics of DNA methylation distribution patterns

by the 5mdC immunolocalization approach presented here

revealed that a decrease in the DNA methylation and its nuclear

redistribution is associated with microspore reprogramming and

embryogenesis initiation, whereas a progressive increase in DNA

methylation accompanies the progression of microspore embryo-

genesis and embryo differentiation [

5 , 7 , 13 ].

The processing of the plant culture sample previous to the

5mdC immunolocalization constitutes a key step which should ﬁ t

with the compromise of preserving the antigenicity together with

the good structural preservation. The processing methods are

Pilar S. Testillano and María Carmen Risueño

493

different for different culture samples due to the different charac-

teristics for in situ cellular analysis on section of the samples, e.g.,

hardness, heterogeneity, cell accessibility, tissue compactness, etc.

At advanced stages of microspore embryogenesis , the individual

developing embryos that can be separated from the microspore or

anther culture s constitute samples with low/mild hardness and

relatively homogeneous structure. Therefore, it could be sectioned

without embedding media by the cryostat, providing thick sections

with good structural preservation. At early stages of the anther

culture, the embryogenic anther is a heterogeneous organ, com-

posed by very different cell types with different wall hardness and

vacuoles. Moreover, the anthers contain microspores and very

early embryos inside the pollen sac which would be lost in non-

embedded sections. Therefore, anthers have to be processed and

embedded in acrylic resins at low temperature, to maintain their

structural integrity and to preserve their antigenic properties as

much as possible. The resins of choice are Technovit 8100 (Heraeus

Kulzer, Wehrheim, Germany) and Lowicryl K4M (Polysciences

Inc, Eppelheim, Germany). In isolated microspore culture s, the

samples of the ﬁ rst stages containing isolated microspores and

small embryos are previously embedded in gelatin in order to

manipulate them as tissue pieces. After that, they are processed like

anthers and embedded in acrylic resins.

The 5mdC immunoﬂ uorescence protocol involves several per-

meabilization steps for thick cryostat sections, including freezing-

thawing, dehydration-rehydration, and mild cell wall enzymatic

digestion. After permeabilization, cryostat sections are treated with

the same protocol than resin sections. Denaturation of the DNA in

sections with HCl is essential to expose the 5mdC antigen to the

antibodies. Further steps included the blocking and the incuba-

tions with the ﬁ rst (anti-5mdC) and secondary (ﬂ uorochrome

Alexa-conjugated) antibodies. The microscopical analysis of the

immunoﬂ uorescence preparations is performed in a confocal laser

scanning microscope (CLSM) which permitted to obtain optical

sections and avoided the out-of-focus ﬂ uorescence of the thick

(30–50 μm) cryostat sections. 1–2 μm semithin resin sections can

be analyzed by both CLSM and epiﬂ uorescence microscopes, even

though the CLSM provided ﬂ uorescent images of higher resolu-

tion and quality (Figs.

1 and 2 ). Controls are performed by elimi-

nating the DNA denaturation by HCl and by immunodepletion

assay s in which the antibody is pre-blocked with the antigen

(5mdC) in vitro, and this pre-blocked antibody is used for immu-

noﬂ uorescence experiments. Negative results of the ﬁ rst control

indicate that the antibody does not cross-react with double-

stranded DNA or other nuclear antigens. Absence of signal in the

immunodepletion experiments indicate that the antibody only rec-

ognized the 5mdC as antigen and did not cross-react with other

antigens, since it was completely blocked in vitro with the 5mdC

(Fig.

1b, c ).

DNA Methylation in Microspore Embryogenesis

494

The method presented here provides unique information on

the DNA methylation nuclear patterns of different plant cell types,

like microspores, pollen grains, anther and embryo cells, and their

dynamics in relation to chromatin organization during prolifera-

tion and differentiation processes that occur during microspore

embryogenesis in different in vitro systems (isolated microspore

Fig. 1 5mdC immunoﬂ uorescence in anthers developed in vivo . Tobacco anthers at the developmental

stage of vacuolated microspore -young bicellular pollen , the most responsive for embryogenesis in vitro induc-

tion. Confocal images of 1 μm semithin Technovit 8100 sections. Same sections of anthers showing micro-

spores (mic) and anther wall (aw) observed with differential interference contrast ( A, B, C ), DAPI staining for

nuclei, cyan ﬂ uorescence ( Aʹ, Bʹ, Cʹ ), and 5mdC immunoﬂ uorescence, green ﬂ uorescence ( Aʺ, Bʺ, Cʺ ). ( A, Aʹ, Aʺ )

5mdC immunoﬂ uorescence. ( B, Bʹ, Bʺ ) Control by immunodepletion of the antibody by in vitro pre-blocking

with 5mdC. ( C, Cʹ, C ʺ) Control by eliminating the DNA denaturation step. The microspore wall, the exine,

showed unspeciﬁ c autoﬂ uorescence of different intensities in cyan and green channels. Bars, 30 μm

Pilar S. Testillano and María Carmen Risueño

495

Fig. 2 5mdC immunoﬂ uorescence during different stages of microspore embryogenesis in different

systems . ( A, Aʹ ) Barley vacuolated microspore before embryogenesis induction. ( B, Bʹ ) Multicellular embryo

still surrounded by the exine (Ex) from an isolated microspore culture of barley . ( C, Cʹ ) Multicellular embryo at

the exine breakdown from an isolated microspore culture of rapeseed. ( D ) Advanced microspore-derived

embryo developed from anther culture of cork oak. ( E ) Advanced microspore-derived embryo developed from

isolated microspore culture of barley. Confocal images of 1 μm semithin Technovit 8100 sections. ( A, B, C )

Differential interference contrast (DIC) images of the same sections than in Aʹ, Bʹ and C ʹ. ( Aʹ, Bʹ, Cʹ, D , E )

Merged images of DAPI staining for nuclei ( blue ) and 5mdC immunoﬂ uorescence ( green ). ( Insets in D , E )

Individual nuclei of advanced microspore embryos of cork oak and barley, respectively. In A ʹ, B ʹ, C ʹ the micro-

spore wall, the exine (Ex), showed unspeciﬁ c autoﬂ uorescence of different intensities in blue and green chan-

nels. Bars in A , A ʹ, 10 μm; in B , B ʹ, C , C ʹ, 20 μm; in D , E , 25 μm

DNA Methylation in Microspore Embryogenesis

496

culture s and anther culture s) and in various plant species, based on

the versatility of the immunolocalization protocol and the good

resolution and quality provided by the CLSM analysis. The infor-

mation raised will give new insights into the mechanisms regulat-

ing epigenetic patterns and chromatin remodeling during in vitro

microspore embryogenesis.

2 Materials

1. Fixative: 4 % paraformaldehyde in PBS. Prepare a solution of 4

% formaldehyde (from paraformaldehyde powder) in PBS, pH

7.0. Heat in a hot bath (no more than 80 °C) until the solu-

tion is transparent. If necessary, a small drop of sodium hydrox-

ide can be added, but pH should be checked afterward. Then

put it on melting ice. Aliquots of freshly prepared 4 % formal-

dehyde solution can be stored at −20 °C and thawed just

before use.

2. Dehydration solutions: Prepare an acetone series of 30, 50, 70,

90, and 100 % in volume of acetone in water, and keep at 4 °C

until use.

3. Resin: Commercial acrylic resin Technovit 8100 (Heraeus

Kulzer, Germany). Prepare the inﬁ ltration solution and embed-

ding solution following the manufacturer instructions, just

before use, and keep them at 4 °C ( see Note 1 ).

1. Cryoprotectant: Prepare a series of sucrose in increasing con-

centrations in PBS, 0.1, 0.5, 1, 1.5, 2, and 2.3 M. Aliquots of

them can be stored at −20 °C and thawed just before use.

2. Commercial OCT (optimal cutting temperature) compound is

kept in liquid-viscous form at room temperature and used to

embed samples during freezing over carbonic ice.

1. Phosphate-buffered saline (PBS).

2. Blocking agents: 5 % BSA and bovine serum albumin (w/v), in

PBS. Dilute 0.5 g BSA in 10 mL PBS with a magnetic stirrer

(without heating or the BSA will coagulate), and prepare

aliquots of 1 mL that can be stored at −20 °C and thawed just

before use.

3. Permeabilization agents: Prepare a methanol series of 30, 50,

70, 90, and 100 % in volume of methanol in water and keep at

room temperature until use. Prepare a mixture of enzymes for

partial cell wall degradation with the following composition:

2.5 % pectinase, 2.5 % cellulase, and 2.5 % pectoliase in

PBS. Prepare aliquots which can be stored at −20 °C and

thawed just before use.

2.1 Fixative,

Dehydration,

and Resin Solutions

2.2 Components

for Processing

in Cryostat

2.3 Solutions for

Immuno ﬂ uorescence

Pilar S. Testillano and María Carmen Risueño

497

4. DNA denaturation agent: Prepare a solution of 2 N HCl (chlor-

hydric acid) in water, and keep it at room temperature until use.

5. Commercial mouse monoclonal anti-5-methyl-deoxy-cytidine

(anti-5mdC) antibody (Eurogentec, Belgium, Cat. Number:

BI-MECY-0100).

6. Commercial goat anti-mouse IgG conjugated to Alexa Fluor

466 secondary antibody (Molecular Probes, Leiden, The

Netherlands).

7. DNA staining agent: Prepare a 1 mg/mL DAPI

(4′,6- diamidino-2-phenylindole) solution in PBS and keep at

4 °C until use.

3 Methods

Two types of sections can be used for in situ analysis of DNA meth-

ylation patterns by 5mdC immunoﬂ uorescence : cryostat sections

and resin sections.

1. Anther and microspore culture samples collected at different

culture times are ﬁ xed overnight with 4 % paraformaldehyde in

PBS at 4 °C. Samples immersed in the ﬁ xative are subjected to

a short vacuum step (1–5 min) for proper penetration of the

ﬁ xative into the cells.

2. After ﬁ xation, samples are washed three times in PBS for 5 min

each washing step.

3. Culture samples of the ﬁ rst developmental stages containing

isolated vacuolated microspores and early multicellular

embryos have to be previously embedded in 15 % gelatin in

PBS and gel solidiﬁ ed on ice for further manipulation, like

embryo or anther samples.

4. Fixed samples can be either dehydrated and resin embedded,

or processed for freezing and cryostat sectioning. The samples

of early stages, which were embedded in gelatin, are dehy-

drated and resin embedded. Larger samples that were not

embedded in gelatin, like anthers, globular, torpedo, and coty-

ledonary embryos, can be processed either for cryostat or resin

embedding.

1. To obtain cryostat sections, ﬁ xed samples are washed in PBS,

and cryoprotected through a gradual inﬁ ltration in sucrose

solutions: 0.1, 0.5, 1, 1.5, and 2 M for 1 h each and 2.3 M

overnight, at 4 °C, embedded in Tissue-Tek optimal cutting

temperature (OCT) compound and frozen on dry ice forming

small pieces of solidiﬁ ed frozen OCT containing the samples at

their interior that should be kept at −20 °C until use.

3.1 Sample

Processing, Section

Preparation,

and Storage

3.1.1 Cryostat Sections

DNA Methylation in Microspore Embryogenesis

498

2. Frozen samples are placed in the cryostat and sectioned at

20–30 μM thickness under −20 °C/−30 °C working

temperature.

3. Cryostat sections are collected on glass slides, washed with

water to eliminate the OCT, and transferred to a water drop

over silanized slides, air-dried and stored at −20 °C until use

for immunoﬂ uorescence ( see Note 2 ).

4. Cryostat sections are then subjected to permeabilization before

their use for immunoﬂ uorescence assays.

1. Fixed samples are dehydrated in an acetone series of 30, 50,

70, 90, and 100 % and then immersed in the Technovit 8100

resin inﬁ ltration solution overnight at 4 °C.

2. After inﬁ ltration, individual samples are embedded in resin

embedding solution ( see Note 3 ) in gelatin capsules which are

covered by a gelatin cap to avoid oxygen that interferes with

the polymerization.

3. Resin capsules are polymerized at 4 °C overnight, and sections

of 1–2 μm thickness are obtained in an ultramicrotome, placed

in a water drop on silanized slides, dried, and stored at 4 °C

until use for immunoﬂ uorescence .

4. Semithin resin sections do not require permeabilization and

are subjected directly to the immunodetection, after incuba-

tion in PBS for a few minutes.

For cryostat sections, permeabilization is required prior to

immunoﬂ uorescence .

1. After thawing the sections at room temperature, they are dehy-

drated and rehydrated in a methanol series (30, 50, 70, 90,

100, 90, 70, 50, 30 %, 5 min each) and PBS.

2. Sections are subsequently subjected to enzymatic digestion of

cell wall

s for additional permeabilization by treatment with an

enzymatic mixture (2.5 % pectinase, 2.5 % cellulase, and 2.5 %

pectoliase) in PBS for 45 min ( see Note 4 ), then washed in

PBS, and subjected to immunoﬂ uorescence (IF) procedure

without drying of the section in any step.

At this step, both section types, cryostat and resin sections, follow

the same protocol.

1. Sections are denatured with 2 N HCl for 45 min, washed in

PB S two times, 5 min each, and then blocked with 5 % (w/v)

bovine serum albumin (BSA) in PBS for 10 min.

2. Sections are then directly incubated for 1 h with the mouse

monoclonal anti-5mdC antibody diluted 1:50 in 1 % BSA in

PBS. After three rinsing steps in PBS, 5 min each, sections are

3.1.2 Resin Sections

3.2 Permeabilization

of Cryostat Sections

3.3 5mdC Immuno-

ﬂ uorescence

on Cryostat and Resin

Sections

Pilar S. Testillano and María Carmen Risueño

499

incubated for 45 min in darkness with the secondary antibody,

an anti-mouse IgG conjugated to Alexa Fluor 488 diluted

1:25 in 1 % BSA.

3. After washing in PBS three times, 5 min each, nuclei are stained

with DAPI (4′,6--diamidino-2-phenylindole) staining solution

for 5 min ( see Note 5 ), washed in sterile water, and mounted

in Mowiol.

4. Immunoﬂ uorescence preparations are then examined under

either an epiﬂ uorescence or a confocal laser scanning micro-

scope (CLSM). CLSM permits to obtain optical sections and

avoid the out-of-focus ﬂ uorescence of the thick (20–30 μm)

cryostat sections. 1–2 μm semithin resin sections can be ana-

lyzed by both CLSM and epiﬂ uorescence microscopes, even

though the CLSM provides ﬂ uorescent images of higher reso-

lution and quality. The results obtained are similar in both

cryostat and resin sections, intense immunoﬂ uorescence sig-

nals on deﬁ ned regions of the nuclei, which are clearly identi-

ﬁ ed by DAPI staining. Confocal optical sections are collected

either at 0.5 or 0.1 μm length intervals in the z axis (section

thickness) for cryostat or resin sections, respectively, and

images of maximum projections can be obtained with software

running in conjunction with the confocal microscope.

Apart from the general control experiments in immunoﬂ uores-

cence assays by eliminating the ﬁ rst and secondary antibodies, two

main controls should be performed to assess the speciﬁ city of the

5mdC immunoﬂ uorescence signal, a control by the elimination of

the DNA denaturation step and another by the immunodepletion

of the 5mdC antibody with the antigen.

This control is performed in samples by applying the whole immu-

noﬂ uorescence protocol and eliminating the DNA denaturation

step by avoiding the HCl treatment before the antibody incuba-

tion. Instead of it, a washing step with PBS during the same time

than the HCl treatment is carried out. The results of this control

should be negative, showing a complete absence of signal which

indicates that the antibody do not cross-react with double- stranded

DNA or other nucleic acid antigens.

1. The anti-5mdC antibodies are pre-blocked with its corre-

sponding immunogen, the 5mdC, by incubating the antibody

in an Eppendorf tube with a 5mdC solution (5 μg/μL in

water) in a proportion of 1:2, v/v, at 4 °C, overnight. During

this time, the immunoglobulins contained in the antibody

solution that speciﬁ cally bind to 5mdC are blocked by the

excess of immunogen in the immunodepletion solution and

cannot bind to any antigen present in the section, providing

negative immunoﬂ uorescence results.

3.4 Controls

for 5mdC Immunoﬂ uo-

rescence Experiments

3.4.1 Control

by Eliminating the DNA

Denaturation Step

3.4.2 Control by

Immunodepletion

of the 5mdC Antibody

with the Antigen

DNA Methylation in Microspore Embryogenesis

500

2. After the above reaction, the pre-blocked antibody solution is

used as primary antibody for immunoﬂ uorescence on the sec-

tions, following the same protocol and conditions as described

above. Negative results of the immunodepletion control exper-

iment indicate that the antibody only recognizes the 5mdC as

antigen and does not cross-react with other antigens in the

sections, since it was completely blocked in vitro with the

5mdC molecules.

The analysis of the immunoﬂ uorescence assays by confocal micros-

copy using the same laser excitation and sample emission capture

settings for image acquisition in all immunoﬂ uorescence prepara-

tions allows the accurate comparison among signals from cells at

different developmental stages and the further quantiﬁ cation of

the signal intensities.

1. For each immunoﬂ uorescence microscopy preparation, confo-

cal optical sections are collected at the same z-intervals, e.g.,

0.5 μm for cryostat sections and 0.1 μm for resin sections, with

the same total number of optical sections (15–20). Then,

images of maximum projections are obtained and used for rela-

tive ﬂ uorescence intensity quantiﬁ cation with software run-

ning in conjunction with the confocal microscope.

2. Fluorescence intensity quantiﬁ cation is performed on random

nuclei of each sample, in a minimum number of nuclei statisti-

cally signiﬁ cant, around 30–50 nuclei per sample (e.g., the

minimum sample size can be estimated by the progressive

mean method).

3. Signiﬁ cant differences among the mean values of relative

5mdC ﬂ uorescence intensities of microspore embryogenesis

developmental stages are compared by appropriate statistical

tests like Student’s t test or one-way variance analysis.

4 Notes

1. Once the inﬁ ltration solution is freshly prepared by mixing the

components of the commercial kit and stored at 4 °C, it can be

used for 1 month either for inﬁ ltration of new samples or to

prepare the embedding solution.

2. In general, for immunoﬂ uorescence of sections, the use of

Teﬂ on- printed multi-well slides (Immuno-Cell Int. Mechelen

Belgium) is very convenient; on one hand, they help to mini-

mize the volumes of antibodies required since the drops of

solutions are conﬁ ned by the well (10 μL, even less in critical

cases, for 7 mm well is enough to cover the section, when

placed in a humid chamber to avoid drying during antibody

3.5 Quantiﬁ cation of

Fluorescence Intensity

in 5mdC Immuno-

localization Confocal

Images

Pilar S. Testillano and María Carmen Risueño

501

incubation). Secondly, they permit to perform individual exper-

iments in each section/well and therefore several assays with

different antibodies, dilutions, or samples in one unique slide.

3. The embedding solution mixture starts its polymerization rap-

idly after its preparation; therefore, it should be prepared in

small quantities to avoid the risk of its polymerization along

the procedure of embedding in capsules. When there are

numerous samples to embed, they can be inﬁ ltrated together,

but the embedding should be performed in consecutive time

steps. For example, prepare only 3 mL of embedding solution

in a tube, proceed with the transfer of the samples to capsules

and ﬁ lling with embedding solution until ﬁ nishing the embed-

ding solution, then prepare another tube with 3 mL, and

repeat with other inﬁ ltrated samples. With this sequential pro-

cedure, the time of manipulation of the embedding solution is

short and it will keep liquid.

4. In case of samples with thick or differentiated cell wall s, or in

the case of thicker sections, the permeabilization should be

more efﬁ cient. In this case, the activity of the enzymes can be

optimized by performing the incubation at higher temperature

(e.g., 37 °C) in a humid chamber or prolonging the time.

5. For thick sections, to facilitate DAPI to penetrate into the

nuclei, add to the staining solution 0.1 % Triton X-100.

Acknowledgments

This work is supported by projects funded by the Spanish Ministry

of Economy and Competitiveness, MINECO, and the European

Regional Development Fund (ERDF/FEDER) of the European

Commission (BFU2008-00203, BFU2011-23752, AGL2014-

52028- R) and Spanish National Research Council, CSIC (PIE

201020E038).

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embryogenesis in Brassica napus . Cytogenet

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gene and protein expression during microspore

embryogenesis by Confocal Laser Scanning

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Touraev A (eds) Advances in haploid produc-

tion in higher plants. Springer Science and

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Coronado MJ, Risueño MC, Testillano PS

(2012) NO, ROS, and cell death associated

with caspase-like activity increase in stress-

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Becerra J, Santos-Ruíz L (eds) Hot topics in

cell biology. Biohealthcare Publishing Limited,

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12. Testillano PS, Solís MT, Risueño MC (2013)

The 5-methyl-deoxy-cytidine localization to

reveal in situ the dynamics of DNA methyla-

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organ and tissue cells during development.

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Pilar S. Testillano and María Carmen Risueño

503

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 29

Embryogenesis and Plant Regeneration from Isolated

Wheat Zygotes

Jochen Kumlehn

Abstract

Wheat zygotes can be mechanically isolated and cultivated to continue their development in vitro. Since

each zygote needs to be individually isolated, only relatively few of these cells are available per experiment.

To facilitate embryonic growth despite of this limitation, the zygotes are kept within a culture insert placed

in a larger dish which itself contains embryogenic pollen cocultivated for continuous medium condition-

ing. This setup ensures that the two cultures, while being physically separated from one another, can

exchange essential intercellular signal molecules passing through the bottom of the insert which is made

of a permeable membrane. Thanks to the natural fate of zygotes, which is to form an embryo followed by

the generation of a plant, embryogenesis and plant regeneration are achieved at much higher efﬁ ciency as

compared to other single-cell systems. While the method is largely independent of the genotype, it allows

for the nondestructive observation, manipulation, and individual analysis of zygotes and very young

embryos.

Key words Cocultivation , Embryonic development , Fertilized egg cell , Single-cell culture

1 Introduction

The commencement of ontogenesis is a fundamental process in

plant development. However, zygotes and young embryos are

hardly accessible to observation over time, manipulation, and cell-

speciﬁ c analyses, because they are hidden under several layers of

tissue within the pistil. In vitro embryogenesis and plant regenera-

tion via culture of isolated zygotes have been preferentially

achieved in Poaceae species such as barley , maize, wheat , and rice

[

1 – 4 ]. Historically, however, zygotes of maize and wheat were

ﬁ rst cultivated following isolation of gametes (egg and sperm

cells) and in vitro fertilization [

5 , 6 ], which was later achieved in

rice as well [

7 ].

The isolation of zygotes and their further embryonic develop-

ment in vitro have been an important technical advance toward the

elucidation of structural patterns and molecular mechanisms in the

504

context of fertilization and early embryogenesis [ 8 , 9 ]. Live-cell

imaging of cultivated wheat and rice zygotes has resulted in valu-

able descriptive information on early embryonic development [

8 ,

10 ]. Isolated zygotes or bicellular proembryos also proved to be

very useful for precise transcriptomic analyses as was shown in

maize [

11 ], wheat [ 9 , 12 ], tobacco , and rice [ 13 , 14 ]. In addition,

isolated barley zygotes were used for stable transgenesis by means

of microinjection of plasmid DNA [

15 ].

Survival and development of plant cells are dependent upon

intercellular exchange of signals, which is typically provided in cell

and tissue culture systems by a suitable cell population density or

sufﬁ cient explant size. To cope with the limited cell number avail-

able per experiment, Kumlehn and coworkers [

16 ] transplanted

isolated wheat zygotes into cultivated wheat or barley ovules,

which succeeded in efﬁ cient embryonic development and plant

regeneration. The cocultivation of heterologous cell types proved

to be a viable alternative approach to effective medium condition-

ing. In the method described here, barley microspores previously

treated to undergo pollen embryogenesis were used for cocultiva-

tion to facilitate embryonic growth of isolated wheat zygotes. To

prevent the zygotes and zygotic embryo s from getting lost in the

comparatively huge population of pollen-derived embryogenic

structures, culture inserts featuring a permeable membrane instead

of a solid bottom are used. Such insert harboring some zygotes is

placed in a larger dish that itself contains the embryogenic pollen

culture so that extracellular signal molecules are allowed to diffuse

from the outer medium portion through the membrane into the

insert, while the zygote- and pollen-derived structures are kept

separated from one another.

2 Materials

All solutions and media are prepared using doubled-distilled water

or equivalent quality and analytical grade chemicals, unless speci-

ﬁ ed otherwise.

1. The German wheat ( Triticum aestivum L.) cultivars Florida

(winter type), Ralle and Remus (spring type) as well as the

Mexican breeding line Veery #5 (spring type) were used to

isolate and cultivate zygotes.

2. Embryogenic pollen cultures used for co-culture with isolated

zygotes were produced in the German barley ( Hordeum vul-

gare L.) cv Igri (winter type).

2.1 Plant Material

Jochen Kumlehn

505

1. Filter paper disks, 7 cm diameter, ash-free, autoclaved.

2. Refrigerated centrifuge equipped with swing-out baskets.

3. Waring blender (Eberbach, MI, USA), sterilizable by heat,

with drive unit.

4. Sterile screw-cap polypropylene centrifuge tubes, 50 mL.

5. Sterile screw-cap round-bottomed polycarbonate cell culture

tubes, 12 mL.

6. Clear-transparent (e.g., Magenta) boxes, ca. 250 mL,

autoclavable.

7. Nylon mesh, 100 μm grid, autoclavable.

8. Hemocytometer, type Rosenthal.

1. Inverted microscope equipped with long-distance condenser

lens, allowing to conduct preparations in culture dishes placed

on the microscope stage.

2. Fine-tipped glass needles, custom- or self-made by a pulling

device.

3. Glass capillary, 100 μM interior diameter, custom- or self-made

by a pulling device.

4. Cell Tram Vario (Eppendorf, Germany) equipped with poly-

propylene tubing.

5. Millicell inserts, 0.4 μm pore size membrane (Millipore,

Germany).

6. 4-well plates, 1.9 cm

2 culture area per well (Nunc, Denmark).

1. K macro minerals [

17 ] (×20): 40.4 g/L KNO

3 , 1.6 g/L

NH

4 NO

3 , 6.8 g/L KH

2 PO

4 , 8.8 g/L CaCl

2 ·2H

2 O, 4.9 g/L

MgSO

4 ·7H

2 O ( see Note 1 ); ﬁ lter-sterilized, stored at room

temperature.

2. K micro minerals [

17 ] (×1000): 8.4 g/L MnSO

4 ·H

2 O, 7.2 g/L

ZnSO

4 ·7H

2 O, 3.1 g/L H

3 BO

3 , 120 mg/L Na

2 MoO

4 ·2H

2 O,

24 mg/L CoCl

2 ·6H

2 O, 25 mg/L CuSO

4 ·5H

2 O, 170 mg/L KI

( see Note 1 ); ﬁ lter-sterilized, stored at 4 °C.

3. Chu N6 macro minerals [

18 ] (×10): 28.3 g/L KNO

3 , 4.62

g/L (NH

4 )

2 SO

4 , 4 g/L KH

2 PO

4 , 1.86 g/L MgSO

4 ·7H

2 O,

1.66 g/L CaCl

2 ·2H

2 O ( see Note 1 ); ﬁ lter-sterilized, stored at

room temperature.

4. Chu N6 micro minerals (×1000): 4 g/L MnSO

4 ·4H

2 O, 500

mg/L H

3 BO

3 , 500 mg/L ZnSO

4 ·7H

2 O, 25 mg/L

Na

2 MoO

4 ·2H

2 O, 25 mg/L CuSO

4 ·5H

2 O, 25 mg/L

CoCl

2 ·6H

2 O ( see Note 1 ); ﬁ lter-sterilized, stored at room

temperature.

2.2 Speciﬁ c

Laboratory Equipment

2.2.1 For the Production

of Barley Embryogenic

Pollen Cultures

2.2.2 For Isolation

and Culture of Wheat

Zygotes

2.3 Stock Solutions

Wheat Zygote Culture

506

5. NaFeEDTA (Ferric sodium ethylenediaminetetraacetate;

75 mM): 2.75 g dissolved in 100 mL; ﬁ lter-sterilized, stored

at 4 °C.

6. CaCl

2 (1 M): 14.7 g CaCl

2 ·2H

2 O dissolved in 100 mL; ﬁ lter-

sterilized, stored at room temperature.

7. KM organics [

19 ] (×100, Sigma K-3129): 2 mg/L

p- aminobenzoic acid, 200 mg/L

L - ascorbic acid , 1 mg/L

D-BIOTIN , 100 mg/L D -calcium pantothenate, 2 mg/L cyano-

cobalamin, 40 mg/L folic acid, 10 g/L myo-inositol , 100

mg/L nicotinamide, 100 mg/L pyridoxine·HCl, 1 mg/L reti-

nol, 20 mg/L riboﬂ avin, 100 mg/L thiamine ·HCl; stored at

−20 °C.

8. Gamborg B5 organics [

20 ] (×1000): 100 mg/L myo-inositol ,

1 mg/L nicotinic acid , 1 mg/L pyridoxine·HCl, 10 mg/L

thiamine ·HCl; ﬁ lter-sterilized, stored at −20 °C.

9. L-glutamine (0.25 M): 1.83 g dissolved in 50 mL with a few

drops of 0.1 M KOH by heating in a water bath; ﬁ lter- sterilized,

stored at −20 °C.

10. Casein hydrolysate (0.1 g/mL, Sigma A-2427): 1 g dissolved

in 10 mL; ﬁ lter-sterilized, stored at −20 °C.

11. Maltose (1 M, ≥99 %): 360 g maltose ·H

2 O dissolved in 1 L;

ﬁ lter-sterilized, stored at room temperature.

12. Maltose (1 M, ≥95 %): 360 g maltose ·H

2 O dissolved in 1 L;

ﬁ lter-sterilized, stored at room temperature.

13. Maltose (0.55 M, ≥95 %): 198 g maltose ·H

2 O dissolved in

1 L; ﬁ lter-sterilized, stored at 4 °C.

14. Glucose (1 M): 180 g dissolved in 1 L; autoclaved, stored at

4 °C.

15. Mannitol (0.4 M): 72.9 g dissolved in 1 L; autoclaved, stored

at 4 °C.

16. Mannitol (0.55 M): 100.2 g dissolved in 1 L; autoclaved,

stored at 4 °C.

17. Xylose (×1000): 1.5 g dissolved in 10 mL; ﬁ lter-sterilized,

stored at 4 °C.

18. IBA (3-indolbutyric acid, 1 mM): 2 mg dissolved in a few

drops of 50 % ethanol, made up to ﬁ nal volume of 10 mL with

hot water ( see Note 2 ); ﬁ lter-sterilized, stored at 4 °C.

19. 2,4-D (2,4-dicholophenoxyacetic acid, 1 mM): 2.2 mg dis-

solved in a few drops of 50 % ethanol, made up to ﬁ nal volume

of 10 mL with hot water ( see Note 2 ); ﬁ lter-sterilized, stored

at 4 °C.

20. BAP (6-benzylaminopurine, 1 mM): 224 mg/L dissolved in a

few drops of 1 M NaOH, made up to ﬁ nal volume of 50 mL

with hot water ( see Note 2 ); ﬁ lter-sterilized, stored at 4 °C.

Jochen Kumlehn

507

21. Kinetin (1 mM): 10.8 mg/L diluted in a few drops of 1 M

NaOH, made up to ﬁ nal volume of 50 mL with hot water ( see

Note 2 ); ﬁ lter-sterilized, stored at 4 °C.

22. Phytagel (×2): 3.5 g suspended in 250 mL cold water; auto-

claved, stored at room temperature.

23. NaOCl (sodium hypochlorite, 2.5 %): 10 mL concentrated

NaOCl (25 %, containing 12 % Cl) diluted in 90 mL water

with three drops of Tween 20; freshly prepared before use.

24. Double distilled water: autoclaved, stored at room

temperature.

25. Tap water: autoclaved, stored at room temperature.

1. Barley pollen culture (KBP,

17 ) medium: 50 mL/L K macro

minerals, 1 mL/L K micro minerals, 1 mL/L NaFeEDTA,

12 mL/L

L -glutamine stock, 10 mL/L KM organics, 4 mL/L

BAP stock, 250 mL/L maltose (1 M, ≥99 %), pH adjusted to

5.9; stored at 4 °C.

2. Zygote culture (N6Z) medium [

3 ]: 50 mL/L Chu N6 macro

minerals ( see Note 3 ), 1 mL/L Chu N6 micro minerals,

10 mL/L KM organics, 27 mL/L

L -glutamine stock,

2.5 mL/L casein hydrolysate stock, 472 mL/L glucose (1 M;

see Note 4 ), 1 mL/L xylose stock, 0.9 mL/L 2,4-D stock, pH

adjusted to 5.7; stored at 4 °C ( see Note 5 ).

3. Regeneration (N6D) medium [

3 ]: 100 mL/L Chu N6 macro

minerals, 2.44 mL/L CaCl

2 stock ( see Note 6 ), 1 mL/L Chu

N6 micro minerals, 10 mL/L KM organics, 20.3 mL/L

L - glutamine stock, 2.5 mL/L casein hydrolysate stock, 1 mL/L

xylose stock, 150 mL/L maltose (1 M, ≥95 %), 2.5 mL/L

IBA stock, 2.3 mL/L kinetin stock, all components mixed in

half of the ﬁ nal medium volume, pH adjusted to 5.7, heated to

about 40 °C, then mixed 1:1 with Phytagel stock previously

melted by heating ( see Note 5 ).

3 Methods

All procedures are carried out at room temperature unless speciﬁ ed

otherwise.

1. Barley or wheat grain is germinated in trays ﬁ lled with 3:1:2

substrate of garden mulch, sand and peat (Substrate 2,

Klasmann, Germany) and placed for 2 weeks in a chamber

providing a 12 h photoperiod (136 μmol/m

2 /s photon ﬂ ux

density) and 14/12 °C (day/night).

2. Seedlings of cvs. Igri ( barley ) and Florida ( wheat ) need to be

vernalized at 4 °C under an 8 h photoperiod for 8 weeks ( see

Note 7 ).

2.4 Nutrient Media

3.1 Growth

of Donor Plants

Wheat Zygote Culture

508

3. Seedlings are transferred to 18 cm diameter pots, ﬁ lled with

2:2:1 substrate formulation of compost, substrate 2 (Klasmann,

Germany), and sand, fertilized by providing 15 g Osmocote

(Scotts Celaﬂ or, Germany; 19 % N, 6 % P, and 12 % K) per pot,

and further held in a chamber providing a 12 h photoperiod

(136 μmol/m

2 /s photon ﬂ ux density) and 14/12 °C (day/

night).

4. As of the tiller elongation stage, the plants are held in a glass-

house at 18/14 °C (day/night) with a minimum of 16 h pho-

toperiod (170 μmol/m

2 /s photon ﬂ ux density) provided by

SON-T-Agro lamps (Philips, Netherlands, ca. 200 W/m

2 )

used in addition to natural daylight if required.

1. Barley spikes are harvested when the tips of the awns have

emerged from the boot. The anthers of these spikes predomi-

nantly contain highly vacuolated, pre-mitotic microspores.

2. The boots are cut and surface-sterilized by spraying with 70 %

ethanol ( see Note 8 ). The ﬂ ag leaf sheath is removed and ﬁ ve

dissected spikes placed onto a moistened, 7 cm ﬁ lter paper disk

per 10 cm Petri dish. After sealing, the plates are held in the

dark at 4 °C for 3–5 weeks.

3. Fifteen pretreated spikes are chopped into ca. 1 cm fragments

and macerated in a Waring blender in the presence of 20 mL

0.4 M mannitol ( see Note 9 ). The blender drive unit is set on

“low” speed and run twice for 15 s.

4. The macerate is ﬁ ltered through a 100 μm mesh into a trans-

parent box. The blender is ﬂ ushed with 10 mL of 0.4 M man-

nitol , which is then also passed through the mesh.

5. The debris remaining on the mesh is squeezed gently to release

further suspension into the box, then returned to the blender

for re-maceration (twice for 10 s) in another 10 mL of 0.4 M

mannitol and the macerate passed through the mesh, which is

again followed by ﬂ ushing the blender.

6. The suspension collected in the box is transferred into a 50 mL

tube, and the box ﬂ ushed with 5 mL 0.4 M mannitol , which is

then added to the tube. The suspension is centrifuged (100 × g ,

10 min, 4 °C).

7. The pellet is re-suspended in 3 mL 0.55 M maltose in a round-

bottomed 12 mL tube with a screw cap. The centrifuge tube is

ﬂ ushed with 2 mL 0.4 M mannitol , which is poured carefully

over the top of the 0.55 M maltose suspension, thereby form-

ing two distinct liquid layers with different density.

8.

The suspension is subjected to density gradient centrifugation

in swing-out baskets (100 × g , 10 min, 4 °C) with the centri-

fuge set to give slow acceleration and deceleration to prevent

the two established layers with different density from becoming

3.2 Production

of Embryogenic Pollen

Cultures of Barley

Used for Cocultivation

Jochen Kumlehn

509

mixed. The interphase, where viable immature, highly

vacuolated pollen have accumulated, is withdrawn by pipet-

ting, transferred to a fresh 50 mL tube to which 10 mL 0.4 M

mannitol is added.

9. The pollen is gently suspended evenly, and a representative

100 μL aliquot is removed to a hemocytometer cell in order to

estimate the population density. Meanwhile the remaining

microspores are pelleted by centrifugation (100 × g , 10 min, 4

°C). Before the supernatant is withdrawn, the tube is left stand

for ca. 5 min to allow still ﬂ oating pollen to settle down.

10. The pellet is re-suspended in an appropriate volume of KBP

medium to deliver a density of 100,000 immature pollen per

1-mL aliquots that are transferred to 35-cm Petri dishes which

are then sealed and incubated until use for cocultivation at 24

°C in the dark ( see Note 10 ).

1. Spikes are manually emasculated 1–3 days before anthesis,

using only the spikelets of the central third of the rachis and

the two major (outer) ﬂ orets per spikelet. All other spikelets

and ﬂ orets are removed from the rachis before detaching the

anthers from the ﬂ orets using ﬁ ne-tipped forceps. To avoid any

unwanted pollination, the spikes are covered by polyethylene

bags.

2. During the period of pistil receptivity, one or two freshly

dehisced anthers taken from non-emasculated spikes are trans-

ferred into each ﬂ oret, so that fresh pollen is released onto the

stigmas.

1. 1–9 h after manual pollination, the spikes are cut, and, after

removal of bracts and lemmas, surface-sterilized in 2.5 %

NaOCl solution for 10 min, then rinsed four times using auto-

claved tap water. All following steps are conducted under asep-

tic conditions using surface-sterile materials.

2. The preparation of tissue is conducted using sterile, ﬁ ne-tipped

forceps and a scalpel ( see Note 11 ). The pistils are carefully

detached from the ﬂ orets and collected in a 35-cm Petri dish,

containing 2 mL of 0.55 M mannitol .

3.

Using a binocular, the lodicules and ﬁ laments are removed

from the pistils, the basal tips are cut without squashing the

tissue (Fig.

1a ) and transferred to another dish, containing 2

mL of 0.55 M mannitol .

4. After having collected some pistil tips with their cut side facing

the liquid surface, the explants are submerged into the solution

to allow them to settle at the bottom of the dish.

5. Using an inverted microscope, the ovule tips are isolated from

the pistil tips, and the remaining outer integument and peri-

carp tissue is discarded (Fig.

1b , see Note 12 ).

3.3 Emasculation

and Manual Pollination

of Florets Used

for Zygote Isolation

3.4 Isolation

of Zygotes

Wheat Zygote Culture

Fig. 1 Isolation and culture of wheat zygotes. ( a ) Cut side of a pistil tip showing pericarp (pc), vascular bundle

(vb), chlorophyll layer (cl), outer integument (oi), and inner integument (ii) (binocular); ( b ) ovule tip consisting of

511

6. Using two ﬁ ne-tipped glass needles, the zygotes can be

gently pushed to be released from the ovule tips (Fig.

1b , see

Note 13 ).

7. Isolated zygotes are collected close to one another at the bot-

tom of the dish (Fig.

1c ) using a glass capillary connected by

polypropylene tubing, ﬁ lled with 0.55 M mannitol solution, to

a manually controlled cell tram. This equipment facilitates to

take up and release single cells in a few nanoliters of liquid.

Before ﬁ lling the system with mannitol solution, the interior of

cell tram, tubing, and glass capillary is to be surface-sterilized

using 70 % ethanol, followed by ﬂ ushing with water.

1. Using the glass capillary, as many as ten zygotes are transferred

onto the membrane of a culture insert containing 100 μL of

N6Z medium and placed in a well of a 4-well plate with the

well containing another 0.35 mL of the same medium.

2. Per well, 0.15 mL of 1–2 weeks old embryogenic pollen culture

is added to the medium outside the culture inserts (Fig.

1d, e ).

After being sealed, the 4-well plates are incubated in a larger

plastic box at 26 °C in the dark for 4 weeks ( see Note 14 ).

3. Macroscopically visible zygote-derived embryos are transferred

to plates containing N6D medium and grown until plantlet

formation (Fig.

1e, f ). Embryos and small plantlets are subcul-

tivated to plates or containers with fresh N6D medium after 3

weeks ( see Note 15 ).

1. Regenerants are transferred to 6 cm diameter pots, ﬁ lled with

Petuniensubstrat (Klasmann, Germany), and placed in a tray

covered by a transparent hood to maintain a high humidity

environment. The tray is placed in a chamber providing a 12 h

photoperiod (136 μmol/m

2 /s photon ﬂ ux density) and

14/12 °C (day/night).

2. After 2 weeks, the hood is removed and the tray left uncovered

for another week.

3. Plantlets are further grown as described above for the donor

plants.

3.5 Zygotic

Embryogenesis

and In Vitro Plant

Regeneration

3.6 Establishment

of Plantlets in Soil

Fig. 1 (continued) inner integument (ii) and displaying degenerated synergid (sy) and zygote (zg) (inverted

microscope); ( c ) Freshly isolated zygotes collected at the bottom of the Petri dish used for dissection (inverted

microscope); ( d ) globular zygotic embryo with suspensor (sp) and embryo proper (ep) residing on the mem-

brane (mb) of a culture insert (ci) after 2 weeks of culture; embryogenic structures (es) derived from coculti-

vated barley pollen are visible behind the membrane (inverted microscope); ( e ) zygotic embryos in a culture

insert (ci) after 4 weeks of culture, embryogenic structures (es) derived from cocultivated barley pollen are

visible in the outer medium portion between insert and culture well (cw) (binocular); ( f ) zygotic embryo on

regeneration medium with primary root (pr), degenerated suspensor (sp), scutellum (sc), and coleoptile (co)

(binocular)

Wheat Zygote Culture

512

4 Notes

1. Components of mineral stocks are dissolved separately before

mixing, and then the whole solution is made up to the required

volume.

2. The use of hot water is to prevent the dissolved molecules from

re-precipitating when being exposed to reduced solvent con-

centration. As soon as the ﬁ nal concentration of the stock is

established, precipitation will no longer appear to happen even

after the solution is cooled down.

3. N6Z contains only half the concentration of macro minerals as

compared to the original N6 medium according to Chu and

coworkers [

18 ].

4. The reason for the use of glucose as major osmoticum and

carbohydrate source in N6Z medium is that a disaccharide,

such as sucrose or maltose , would effect a higher speciﬁ c den-

sity which entails freshly isolated zygotes to ﬂ oat to the medium

surface where most of them would burst.

5. Since the addition of some organic acids formerly used in N6Z

and N6D media [

3 ] proved to be unnecessary, these can be

omitted.

6. N6 minerals [

18 ] need to be supplemented with additional

CaCl

2 to obtain a ﬁ nal concentration of 3 mM, so making sure

that the medium will solidify using Phytagel .

7. The vernalization treatment is not required in the spring-type

accessions.

8. All steps following the surface-sterilization of the boots are to

be conducted under aseptic conditions, using surface-sterile

equipment and solutions.

9. All equipment and solutions used to process the pretreated

spikes and immature pollen need to be precooled to 4 °C and

should be kept on ice.

10. 1–2 week-old embryogenic pollen cultures are used for cocul-

tivation with isolated zygotes, whereas cultures older than 2

weeks showed a reduced capability of supporting zygotic

embryo genesis in vitro.

11. To keep the scalpel blade and forceps clean, it is advisable to

remove tissue debris after each preparation step using a piece

of household viscose foam, autoclaved and moistened with dis-

tilled water.

12. While the zygote is visible through the inner integument that

forms the isolated ovule tip, the removal of the outer integu-

ment is essential to facilitate zygote isolation.

Jochen Kumlehn

513

13. Wheat zygotes are shaped pear-like as long as being embedded

in ovular tissue. Owing to the step-wise release from the stabi-

lizing embryo sac and the use of hypotonic solution for the

isolation procedure, the zygotes take on a spherical form before

they can be recognized within the ovule tips (Fig.

1b ). Since

the zygote is protoplast-like and not interconnected with

neighbor cells via plasmodesmata, it can be mechanically iso-

lated without the use of cell wall -degrading enzymes. The

diameter of isolated wheat zygotes is 60–80 μm.

14. In vitro zygotic embryo genesis f ollows fairly the same pattern

formation as the one of embryos growing in planta . While the

ﬁ rst zygotic cell division appears to be symmetrical with regard

to the volume of daughter cells and is generally completed

within 24 h after pollination, the following period of 1–2 weeks

of bisymmetric pro embryo development includes the differen-

tiation of suspensor and embryo proper (Fig.

1d ). This is fol-

lowed by dorsoventral embryo development, characterized by

the formation of scutellum, shoot apical meristem, coleoptile,

primary root primordium, and coleorhiza.

15. The efﬁ ciency of embryo formation and plant regeneration

proved to be genotype independent. However, a drop in isola-

tion efﬁ ciency was observed when exotic accessions with

smaller grains were used [

21 ]. Using the described procedure,

80–90 % of the isolated zygotes form embryos, most of which

are capable of plant regeneration. The plants obtained show

phenotypically normal development and grain set.

Acknowledgment

I would like to thank my colleague Dr. Maia Gurushidze for criti-

cally reading the manuscript.

References

1. Holm PB, Knudsen S, Mouritzen P, Negri D,

Olsen FL, Roué C (1994) Regeneration of fer-

tile barley plants from mechanically isolated

protoplasts of the fertilized egg cell. Plant Cell

6:531–543

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Jochen Kumlehn

515

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 30

From Somatic Embryo to Synthetic Seed in Citrus

spp. Through the Encapsulation Technology

Maurizio Micheli and Alvaro Standardi

Abstract

In vitro propagation by somatic embryogenesis represents an efﬁ cient alternative method to produce

high- quality and healthy plants in Citrus species. The regenerated somatic embryos need protection from

mechanical damages during manipulation and transport, as well as nutritive support for their evolution in

plantlets after sowing. The encapsulation technology allows to obtain synthetic seeds by covering somatic

embryos with a gel of calcium alginate enriched by nutrients. This chapter describes the procedure for

producing synthetic seeds containing somatic embryos from different Citrus genotypes.

Key words Artiﬁ cial seed , Calcium alginate matrix , Plant tissue culture , Somatic embryo genesis ,

Synseed

1 Introduction

The increasing world’s demand for new and promising Citrus gen-

otypes requires effective and innovative technologies for high-

quality plant production. Consequently, research is looking for an

innovative procedure able to join the advantages of micropropaga-

tion (high productive efﬁ ciency, sanitary plant conditions, and

reduced space requirements) with the technologic characteristics

of the zygotic seed, as handling, storability, and transportability

[

1 ], actually represented by the synthetic seed technology. The

original concept of synthetic seed ( artiﬁ cial seed or synseed ) was

applied to desiccated or hydrated somatic embryo s (SEs) and did

not involve the encapsulation [

2 , 3 ]. Later Murashige [ 4 ] gave the

ﬁ rst deﬁ nition of synthetic seed as “an encapsulated single SE

inside a covering matrix.”

The large use of sodium alginate as encapsulating agent is due

to its moderate viscosity , low spin ability of solution, low toxicity,

quick gellation, low cost, and biocompatibility characteristics [

5 – 7 ].

The encapsulation technology was proposed to safeguard the SEs

from mechanical damages during handling in the nursery and

516

transportation in the farms, as well as to provide nutrients ( artiﬁ cial

endosperm ) during their evolution in plantlets under in vivo or

in vitro conditions ( conversion ). In fact, SEs are structurally similar

to gamic or zygotic embryo s, but lack nutritive and tegument struc-

tures [

5 ]. Nevertheless, the ﬁ rst experiments on the encapsulation

were conducted employing SEs, as their bipolar nature, able to con-

vert in plantlets in a single step, made them suitable for synthetic

seed production [

2 , 8 ]. SEs develop from somatic cells, and this

regenerative pathway allows the clonal propagation. Their use as

encapsulated explants for synthetic seed preparation is however lim-

ited because of the involved difﬁ culties, due to asynchronism dur-

ing SEs formation and development, somaclonal variation , recurrent

embryogenesis [

8 ], and embryo dormancy [ 9 ]. Moreover in vitro

SE production requires expensive manual labor, even though they

could be obtained by bioreactors [

10 ]. Therefore, different propa-

gules were tested to produce synthetic seeds. New perspectives

emerged with the use of non-embryogenic unipolar plant propa-

gules. In fact, the most recent concept involves every meristematic

tissues (in vitro or in vivo derived), as long as able to convert in a

whole plantlet after encapsulation and possible storage [

5 , 7 , 8 ,

11 – 14 ]. However, the abovementioned limitations of SEs for syn-

thetic seeds production seem to be infrequent in Citrus spp., and

several studies are focused on the application of the encapsulation

technology to citrus [

7 , 15 – 21 ].

2 Materials

Since some researchers found that the SE size affects the con-

version in different plant species [

22 – 25 ], we carried out pre-

liminary experiments using different sized SEs of Citrus

genotypes for encapsulation (unpublished data). The results

indicated that the largest SEs (5–6 mm) showed the highest

values in terms of viability (green appearance of explants, with

no necrosis or yellowing), regrowth (increasing in size of the

explants with consequent breakage of the involucre and extru-

sion of at least one visible shoot or root after the sowing ), and

conversion [

16 , 17 ]. Nevertheless their encapsulation involves

the formation of an irregular alginate layer around the propa-

gule, reducing the protective and nutritive functions. So, in our

experiments, we used only medium-sized SEs (3–4 mm) dis-

carding the larger and the smaller ones (Fig.

1 ), hence limiting

the negative effects of asynchronism ( see Note 1 ) and recurrent

embryogenesis ( see Note 2 ).

Usually our experiments were carried out using hydrated SEs of

Citrus reticulata Blanco cv Mandarino Tardivo di Ciaculli, Citrus

limonimedica Lushington, and Citrus clementina Hort. ex Tan. cvs

Nules and Monreal, obtained according the procedures described

2.1 Plant Material

Maurizio Micheli and Alvaro Standardi

517

by Germanà and co-workers [ 26 – 30 ]. The synthetic seeds of these

genotypes were sown and maintained in aseptic conditions. In addi-

tion, synthetic seeds of Citrus reticulata Blanco cv Mandarino

Tardivo di Ciaculli were sown also in non sterile conditions.

1. Tissue culture facilities: Graduate cylinders, pipettes, lab pipet-

tor, glass beakers, magnetic stirrer, spin bar, analytical balance,

lab spoons, weighing boats, pH meter, NaOH and HCl solu-

tion (0.1 N), 100 mL screw capped Pyrex glass jars, autoclave,

horizontal ﬂ ow cabinet, forceps, scalpels, blades, and electric

incinerator.

2a. Aseptic conditions: Distilled water, half strength MS basal

medium [

31 ], 0.25 g/L malt extract , 0.25 g/L ascorbic acid ,

1 mg/L gibberellic acid , ( GA

3 ), 0.02 mg/L a-naphthalene

acetic acid (NAA), and 68 g/L sucrose ( artiﬁ cial endosperm ).

b. Nonsterile conditions: Artiﬁ cial endosperm and 100 mg/L

Thiophanate-methyl TM

® (Pestanal, Riedel-de-Haen).

3. Alginate sodium salt, medium viscosity (2.5 % w/v).

4. Calcium chloride anhydrous (1.1 % w/v).

1. Tissue culture facilities: Graduate cylinders, pipettes, lab

pipettor, glass beakers, magnetic stirrer, spin bar, analytical

balance, lab spoons, weighing boats, pH meter, NaOH and

HCl solution (0.1 N), Magenta

® jars (7 × 7 × 7 cm), autoclave,

horizontal ﬂ ow cabinet, forceps, scalpels, blades, electric

2.2 Encapsulation

Solutions

2.3 Sowing Media

and Culture Conditions

Fig. 1 Synthetic seeds obtained from different sized SEs of Citrus

Synthetic Seed in Citrus spp.

518

incinerator, and growth chamber (temperature of 21 ± 2 °C,

photosynthetic photon ﬂ ux density of 40 μmol/m

2 /s, and

photoperiod 16 h).

2a. Aseptic conditions: Distilled water, full strength MS basal

medium [

31 ], 0.5 g/L malt extract , 0.5 g/L ascorbic acid , 68

g/L sucrose and 7 g/L agar , and ﬁ lter paper bridges.

2b. Nonsterile conditions: Filter paper bridges, perlite, soil

(Compo-Cactea

® ), or Jiffy-7 Pellets (J7).

3 Methods

Three solutions are required to encapsulate SEs: coating , complex-

ing , and rinsing solutions (Fig.

2 ). The common component is

represented by the artiﬁ cial endosperm ( see Subheading

2.2 ) added

of 2.5 g/L sodium alginate ( coating matrix ) and 1.1 g/L calcium

chloride ( complexing solution ). The rinsing solution is composed

only by the artiﬁ cial endosperm . All solutions and media are

adjusted to pH 5.5 and autoclaved at 115 °C for 20 min just after

their transferring into the containers. During the autoclaving cycle,

the sodium alginate is completely dissolved forming a dense dark

yellow solution. The artiﬁ cial endosperm of the synthetic seeds

sown in nonsterile conditions is enriched by Thiophanate-methyl

TM

® ( see Note 3 ).

Fig. 2 Coating, complexing, and rinsing solutions employed for encapsulation of

Citrus SEs ( from left to right )

Maurizio Micheli and Alvaro Standardi

519

1. Single SEs are immersed in alginate solution for a few seconds

( see Note 4 ).

2. The alginate-coated SEs are then dropped into the complexing

solution for 25–30 min ( see Note 5 ).

3. The encapsulated SEs are washed 2–3 times in the rinsing solu-

tion for 10–15 min in order to remove the toxic residual ions

of chloride and sodium ( see Note 6 ). The whole procedure is

carried out in aseptic conditions under a horizontal ﬂ ow

cabinet.

1a. Aseptic conditions: After washing, the synthetic seeds are asep-

tically transferred into closed Magenta

® jars, containing steril-

ized agar sowing medium or ﬁ lter paper bridge, moistened

with 10 mL of artiﬁ cial endosperm ( see Note 7 ).

b. Non-sterile conditions: After washing the synthetic seeds are

aseptically transferred into Magenta

® jars containing sterilized

ﬁ lter paper bridge, perlite, soil (Compo-Cactea

® ), or “Jiffy-7

Pellets” (J7) moistened with appropriate amount of artiﬁ cial

endosperm ( see Note 7 ).

2a. Aseptic conditions: The Magenta

® jars are hermetically closed,

and the cultures are transferred into the growth chamber.

b. Nonsterile conditions: The cultures are then transferred into

the growth chamber, and the Magenta

® jars are not hermeti-

cally closed, allowing the gas exchanges and the water evapora-

tion. To prevent the synthetic seeds dehydration, the substrates

moisture is periodically monitored and restored with distilled

water.

3. After 1 week, fungal or bacterial contamination is monitored.

4. After 45 days, viability, regrowth and conversion (Fig.

3 ) are

evaluated.

4 Notes

1. The asynchronism involves the simultaneous presence of dif-

ferent sized SEs at the end of regenerative cultures. Their

encapsulation determines the formation of heterogenous syn-

thetic seeds with different ability and energy of conversion . So,

the synchronism is a crucial step in taking advantage of somatic

embryo genesis for the commercial production of plants by

synthetic seeds.

2. Recurrent or secondary somatic embryo genesis in the produc-

tion of new SEs from the mature ones.

3. The application of synthetic seeds in the nurseries should imply

their conversion in non-sterile conditions using substrates as

perlite, sand, paper, or peat. In this case, the protection of the

3.1 Encapsulation

3.2 Sowing

and Evaluation

Synthetic Seed in Citrus spp.

520

synthetic seeds from fungal and bacterial contaminations dur-

ing conversion is essential. The beneﬁ cial effect of Thiophanate-

methyl fungicide on the Citrus synthetic seeds conversion has

been showed [

17 ].

4. In substitution to sodium alginate , several substances were

tested, like mixture of sodium alginate with gelatin, potassium

alginate, polyco 2133, carboxymethyl cellulose, carrageenan,

Gelrite , guar gum, sodium pectate, and tragacanth gum

[

5 – 7 ].

5. During the complexation step, ion exchange occurs through

the replacement of Na

+ by Ca

2+ , forming calcium alginate by

ionic cross-linking among the carboxylic acid groups and the

polysaccharide molecules and producing a polymeric structure

called “egg box” [

13 , 32 , 33 ]. Hardening of calcium alginate

bead is affected by the concentration of sodium alginate and

calcium chloride , as well as the complexing time. Usually, at

higher consistence corresponds good protection during trans-

port and manipulation but higher difﬁ culty of explants in

breaking the alginate coat [

8 ].

Fig. 3 Extrusion of shoot and root apex ( black arrows ) from the alginate matrix at

the beginning of conversion

Maurizio Micheli and Alvaro Standardi

521

6. Automation systems have been proposed, as somatic

embryo genesis and encapsulation are expensive techniques

due to the high manual labor requirement. The use of bioreac-

tors for temporary immersion system has shown to be effective

for the production of Citrus deliciosa SEs [

34 ]. Concerning

automation, several devices are available for the encapsulation of

SEs or other in vitro-derived vegetative propagules, using systems

based on concentric tube nozzle, multiple wire loops, rotating

disks, perforated plates, or precision dripping [

10 , 35 , 36 ].

7. Before sowing , the synthetic seeds can be stored at 4–6 °C in

darkness, using closed sterile dishes or vials containing some

drops of artiﬁ cial endosperm solution to avoid dehydration.

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Maurizio Micheli and Alvaro Standardi

523

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 31

From Stress to Embryos: Some of the Problems

for Induction and Maturation of Somatic Embryos

Sergio J. Ochatt and Maria Angeles Revilla

Abstract

Although somatic embryogenesis has been successfully achieved in numerous plant species, little is known

about the mechanism(s) underlying this process. Changes in the balance of growth regulators of the cul-

ture medium, osmolarity, or amino acids as well as the genotype and developmental stage of the tissue used

as initial explant may have a pivotal inﬂ uence on the induction of somatic embryogenic cultures. Moreover,

different stress agents (ethylene, activated charcoal, cold or heat or electrical shocks), as well as abscisic

acid, can also foster the induction or further development of somatic embryos. In the process, cells ﬁ rst

return to a stem cell-like status and then either enter their new program or dye when the stress level

exceeds cell tolerance. Recalcitrance to differentiation of somatic cells into embryos is frequently observed,

and problems such as secondary or recurrent embryogenesis, embryo growth arrest (at the globular stage

or during the transition from torpedo to cotyledonary stage), and development of only the aerial part of

somatic embryos can appear, interfering with normal germination and conversion of embryos to plants.

Some solutions to solve these problems associated to embryogenesis are proposed and two very efﬁ cient

somatic embryogenesis protocols for two model plant species are detailed.

Key words Arabidopsis thaliana , Embryogenesis recalcitrance , Medicago truncatula , Stress agents

1 Introduction

Somatic embryo genesis is in many species the predominant plant

regeneration pathway, during which dedifferentiated somatic plant

cells become totipotent and develop into embryos which, subse-

quently, convert into plants [

1 – 3 ]. Surprisingly, although somatic

embryo genesis is widely used for propagation and has in recent

years become one of the preferred regeneration methods for com-

mercially cultivated biotech crops, little is known about the mecha-

nisms underlying this process. One of the main interesting features

of somatic embryogenesis is that it may sidestep aging limitations

of cultures to yield large numbers of embryos from elite genotypes

including for woody species [

4 ], where somatic embryogenesis is

likely the only way of producing juvenile tissues of recalcitrant

524

species [ 5 ]. In addition, as somatic embryo development resembles

that of zygotic embryo s [

2 , 6 ], they are also an interesting model

system to understand the physiological, biochemical, and molecu-

lar mechanisms taking place during embryo development [

6 – 9 ].

The use of somatic embryogenesis has been reviewed in a large

number of species [

10 – 12 ]. The ability of cells to retain totipo-

tency and developmental plasticity in a differentiated stage makes

them unique and renders them capable to dedifferentiate, prolifer-

ate, and subsequently regenerate into mature plants, provided that

optimum culture conditions are developed [

3 , 13 , 14 ].

Embryogenic cultures are initiated from primary explants on a

medium containing mainly an auxin alone but often also a cytoki-

nin [

1 , 6 , 15 , 16 ] and, sometimes, even only a cytokinin [ 16 ]. The

most commonly used auxin is 2,4-dichlorophenoxyacetic acid

(2,4-D) [

1 , 12 , 15 , 17 ], which has been suggested to downregu-

late gene expression through changes in the level of DNA methyla-

tion [

14 , 18 ]. In addition, various culture conditions and treatments

have an impact on the induction of somatic embryo genesis, includ-

ing balance of plant growth regulator s [

15 , 17 ], medium osmolar-

ity [

19 , 20 ], pH [ 1 ], amino acid, or salt concentration [ 21 ], while

the most inﬂ uential traits identiﬁ ed so far remain the particular

genotype studied and the developmental stage of the tissue used as

initial explant [

22 ].

Somatic embryo genesis may be initiated either directly by

inducing embryos to develop on the surface of the initial explant,

or indirectly, via an intermediary step of callus formation from

which the embryos subsequently regenerate [

6 , 21 ]. Once embryo-

genic cells are initiated, they undergo a continuous, unlimited

cycle producing further pro-embryogenic or embryogenic masses

[

23 ], resulting in multiplication of the original plant. A special case

of somatic embryo genesis is secondary or recurrent embryogene-

sis, which occurs when the ﬁ rst somatic embryo formed fails to

germinate and, instead, gives rise to new successive cycles of

embryogenesis [

17 ]. In some species, this has been sought as a

means of cloning embryogenic lines as the process can be main-

tained indeﬁ nitely [

5 , 24 , 25 ]. However, secondary embryos

develop directly from epidermal and subepidermal cells of embryos

[

26 ], mostly at their root pole or on the main axis and cotyledons

[

27 – 29 ], whereby they interfere with normal germination of the

original embryo and, in ﬁ ne being repetitive, with the conversion

to plants. For somatic embryos, to reach the cotyledonary stage

and then accumulate the storage products needed for conversion

to plants [

6 ], the medium and culture conditions have to be

changed. One of the main growth regulators in embryo matura-

tion in vivo is abscisic acid ( ABA ), and treating embryogenic cul-

tures in vitro with this hormone has been beneﬁ cial in some species

[

6 , 23 , 29 , 30 ], particularly because of its involvement in the

acquisition of partial desiccation [

5 , 25 ] or cold tolerance of mature

Sergio J. Ochatt and Maria Angeles Revilla

525

somatic embryos that precedes their competence to germinate

[

17 , 23 ]. Likewise, ethylene , activated charcoal [ 31 ], pH [ 1 ], cold

[

20 , 32 ] or heat [ 1 ] shocks, osmotic stress [ 19 , 20 , 33 , 34 ], elec-

tricity [

35 ], and even centrifugation [ 36 , 37 ], and sonication

[

37 – 39 ] have been reported to foster somatic embryo induction

but also maturation in different species. No such clear effect can be

ascertained for light conditions. Dark culture has been beneﬁ cial

for embryogenesis, reducing the activity of enzymes responsible

for release of phenols that induce callus and early embryo brown-

ing in species prone to suffering from such phenomena [

3 , 4 , 15 ,

17 ]. On the other hand, photoperiodic light regimes have been

preferred and even required for other species [

37 , 40 ]. Other stress

agents such as heavy metals, starvation, and wounding have also

been reported to promote responses in several models [

41 , 42 ],

and they are moreover an integrating part of dedifferentiation. In

this respect, several studies have shown that, prior to redifferentiat-

ing, cells ﬁ rst return to a stem cell-like status and then either enter

their new program or dye when the stress level exceeds cell toler-

ance or when a mediation of cell responses to stress is hampered by

their physiological status [

3 , 6 , 43 , 44 ]. Indeed, stress-induced

morphogenetic response has been ascribed to the redirection of

growth to better acclimate to an exposure to stress [

13 , 42 , 45 ].

Only those mature somatic embryo s with a normal morphol-

ogy having accumulated enough storage products will be able to

convert into normal plants [

6 , 7 , 29 ]. Following transfer to a ger-

mination medium, somatic embryos develop similarly to zygotic

embryo s, yielding plants that should be true to type [

6 , 16 , 33 ,

46 ]. The hormonal composition needed for germination of the

somatic embryos will mostly depend on the species (and some-

times genotype) studied and there is no generally applicable rule.

Hormone-free media have been reported for both herbaceous [

1 ,

29 ] and woody [ 47 ] species, but media with various auxin/cytoki-

nin contents have been employed with species as wide apart as

legumes [

16 ] and forest trees [ 17 ]. Besides a number of publica-

tions referred the need to add extra miscellaneous compounds to

the medium such as glutamine , casein hydrolysate , etc. [

1 , 17 ],

conversely, there is consensus in the literature on the conditions

required to acclimatize the somatic embryo-derived plantlets to

in vivo conditions which is similar to that usually employed for

micropropagated plants [

1 ].

The most frequent applications of somatic embryo genesis are

the mass propagation of selected material, obtained after in vitro

selection or genetic transformation [

5 , 6 , 17 , 48 , 49 ], and there

are several examples of its commercial exploitation, in particular for

gymnosperms [

23 ], while this is generally still to be done for most

angiosperms with the exception of several ornamentals. It is also

employed for a better understanding of various fundamental mech-

anisms and processes, including those dealing with the acquisition

From Stress to Embryos

526

and eventual loss of regeneration competence, as well as for the

recovery of novel genotypes following in vitro selection for stress

tolerance, somatic hybrid ization, or gene transfer. These aspects

have already been reviewed and discussed in the past. Here, we

shall focus on the problems that may arise during somatic embryo-

genesis and we shall also discuss some possible solutions for them.

2 Development of Somatic Embryos Formed In Vitro

Somatic embryo genesis is one of the two major pathways for plant

regeneration in vitro, and it may take place from undifferentiated

tissues (protoplasts, cell suspension s, callus) but also from highly

differentiated cells (immature gametes), leading respectively to

regeneration of normal plants (that should resemble the mother

plants) or to haploids that will thereafter have to undergo chromo-

some doubling for genome ﬁ xation. Since somatic embryo s can

arise from a single cell, it is a way of choice to regenerate transgenic

plants. The process includes a sequence of developmental stages,

the ﬁ rst of which is often the induction of callus from explants

(Fig.

1a ), followed by the induction of somatic embryos from such

callus tissues that will thereafter follow a common developmental

path from globular- to heart-shaped embryos (Fig.

1b ), then tor-

pedo-shaped embryos (Fig.

1c ), and ﬁ nally mature cotyledonary

embryos (Fig.

1d ) which are capable of “germinating” (Fig. 1e ),

i.e., of converting into whole viable plantlets. To date, these general

Fig. 1 A typical sequence of somatic embryo genesis from root- or leaf-derived callus of Arabidopsis . ( a ) Leaf

explant starting to produce callus. ( b ) A highly embryogenic callus with many somatic embryos at early stages

of development, i.e., from globular to heart. ( c ) A cluster of torpedo stage embryos showing new globular

embryos developing on one of the torpedo embryos ( arrow ). ( d ) A highly embryogenic cluster with mostly

torpedo to cotyledonary stage embryos. ( e ) Liquid culture of embryogenic clusters whereby, with time, only

embryos and plantlets proliferate in culture

Sergio J. Ochatt and Maria Angeles Revilla

527

steps have been successfully applied to many species [ 50 ]. In this

context, auxin is required to induce and maintain a high rate of

proliferation of unorganized plant cells, but low-auxin or simply a

hormone- free medium is needed to induce those developmental

responses that are normally dependent on endogenous hormonal

factors [

43 , 51 ]. Embryos of a unicellular origin are similar to

globular zygotic embryo s and they are sometimes connected to the

maternal tissue by a suspensor like structure, while those derived

from multiple cells initially look like a smooth and bright nodule

where the embryos at its base are usually connected to the mater-

nal tissue through their epidermis [

52 ]

Not all plant cells are capable of expressing totipotency in vitro, a

process that strongly depends on the genetic background, the

physiological status of the donor plant, the type of explant, and its

physiological/developmental status, the culture medium and con-

ditions, and any possible interactions among all these factors

[

15 , 53 ]. Activation of key regulators of somatic embryo genesis is

preceded by a reprogramming of cellular metabolism which is

often induced by some kind of physiological stress and will not be

expressed by somatic cells although it already potentially exists in

the plant genome [

13 , 41 , 42 , 51 ]. Thus, the developmental switch

from a somatic to an embryogenic status in cells occurs under the

inﬂ uence of both physical and chemical inductors but also requires

a major and dynamic reprogramming in gene expression [

7 , 41 ,

42 , 54 ], entailing the activation of a number of signal cascades

leading to a differential (released) gene expression which, in turn,

renders the undifferentiated cells capable to acquire an embryo-

genic capacity [

6 , 13 , 43 ].

As opposed to the highly efﬁ cient embryogenesis sequence

shown in Fig.

1 , there are many species where attaining this is still

difﬁ cult and sometimes even impossible. Indeed, there are three

key stages in this process where a blockage may occur, two of them

2.1 Some of the

Problems Arising

During Somatic

Embryogenesis

and Possible Solutions

Fig. 2 Some problems that may be found during early stages of somatic embryo genesis in pea ( Pisum sativum

L.). ( a ) Browning of globular and heart somatic embryos ( arrows ) and of the callus supporting them. ( b ) Callus

overgrowth and secondary somatic embryogenesis on developing embryos that will fail to convert into plants.

( c ) A cluster of abnormal somatic embryos showing browned roots ( solid arrows ) and fused cotyledons ( dotted

arrow ). ( d ) Pale embryos that have not accumulated storage products are blocked at the late cotyledonary

stage and are unable to germinate

From Stress to Embryos

528

during early development of embryos (Fig. 2 ) and the third one at

the latest stages leading to rooting (Fig.

3 ), as follows:

1. The earlier stages of somatic embryo development, with

embryo growth arrested at the globular stage, whereby mito-

ses stop and embryos start to brown or are covered by de novo

callus overgrowth (Fig.

2a ), which may also sometimes be

associated with secondary embryogenesis (repetitive or not)

(Fig.

2b ).

2. During the transition from torpedo to cotyledonary embryos,

where abnormal (i.e., “trumpet” (Fig.

2c )) fused cotyledons

are formed and/or embryos become pale in color (Fig.

2d ),

due to a lack of accumulation of the storage compounds needed

for the somatic embryo to mature and eventually germinate.

3. At the end of embryo development where, as may happen also

with organogenesis-derived regenerants, only the aerial part of

the somatic embryo s develops (Fig.

3a ) and they have to be

transferred to a different medium for rooting. Then, while the

root pole should grow fast, in an unsuitable medium callus

starts to proliferate instead (Fig.

3b ), mostly at the junction

between the aerial part and the root of the somatic embryo-

derived plantlet (Fig.

3c ). Such plantlets often lack vascular

connection between the root and shoot portions and die upon

transfer in vivo.

Species with a tendency to undergo any one or all of these

processes are regarded as recalcitrant to somatic embryo genesis

and thus to biotechnology approaches based on it for the recov-

ery of novel genotypes, as in haplo-diploidization and various

genetic transformation protocols, e.g., with legumes [

16 , 55 ]

and cereals [

13 ].

As the developmental blockages above tend to concern differ-

ent processes, they require different solutions. It is difﬁ cult to sug-

gest general strategies that will successfully resolve these problems

Fig. 3 Problems with rhizogenesis from somatic embryo s of Medicago truncatula . ( a ) By 9 days on an unsuit-

able medium with root primordia browned and recallusing. ( b ) Embryos of the same age on the right medium.

( c ) Senescent seedling due to the lack of vascular connection between the roots and the aerial part. ( d )

Close-up of hypocotyl/root section of the plantlet in ( c ) showing the interfering callus proliferation

Sergio J. Ochatt and Maria Angeles Revilla

529

but some simple measures can often be applied to at least palliate

or delay their occurrence and thereby permit the recovery of

somatic embryo -derived normal and fertile plants. Thus, for block-

ages occurring at the earliest stages above, shortening the periodic-

ity between subcultures restrains callus development and allows

very immature embryos to develop better. This operation, how-

ever, is not without risks as early globular somatic embryos must be

excised and transferred individually for subsequent development

on a medium that, generally, will have to be enriched in cytokinins

and, sometimes, also in gibberellic acid . This problem is frequently

observed among protoplast-derived callus of pea [

29 ], and several

early cytological predictors of the acquisition of somatic embryo-

genesis competence [

19 ] may help to monitor the evolution of

callus tissues during their culture, in order to act before the shift

from embryogenic back to a sporophytic path is onset. Also at this

stage, increasing the agar concentration of the medium or, like-

wise, increasing its osmolarity either by replacing, at least partially,

the sucrose or glucose with a polyalcohol such as mannitol , or by

adding polyethylene glycol (MW 6000), may also be helpful to

slow down the callus proliferation while having little deleterious

effect on the developing somatic embryos [

15 , 20 , 34 ]. On the

other hand, when the problem encountered is secondary

embryogenesis, a distinction should be made between situations

where this process is repetitive and those where the ﬁ rst secondary

embryos formed will normally convert into plants. Thus, in the

latter case the best solution is probably to simply disregard the

problem and try to enhance secondary embryogenesis instead, so

as to increase the potential for regeneration. Conversely, when the

process is constant and no embryos ever germinate, little informa-

tion about possible solutions is available in the literature. One pos-

sibility that has worked to date with several genotypes of legumes

[

15 ] is to add abscisic acid to the medium for at least one passage

and in combination with an auxin.

For situations when embryo development stops at the transi-

tion from torpedo to cotyledonary stage, modiﬁ cations to the

nutrient composition of the medium may prove appropriate to

warrant a sufﬁ cient accumulation of storage compounds as needed

for embryo maturation . In this respect, it has been shown that

nitrogen, sulfur [

6 ], and also sugar (type and concentration, 7 ) in

the medium play a major role in embryo maturation. Likewise,

some studies have shown that introducing a mild ionic stress (Na

+

or K

+ ) at this transition stage might favor embryo maturation [ 34 ].

Finally, when the main blockage found is for the rooting of shoots

derived from somatic embryo s (whose root pole would not develop

easily on the embryogenic media), this is likely due to the use of an

unsuitable combination of medium salt-strength and hormone

(auxin) content. Such cultures should be treated as for convention-

ally propagated difﬁ cult-to-root shoots, i.e., by testing a reduction

From Stress to Embryos

530

of the salt-strength to half (or even less in very recalcitrant species

such as some cereals and various neglected crops ), by replacing a

strong auxin by a weaker one (i.e., if NAA was used, replacing it

with IBA or IAA), or by totally deleting auxin from the medium.

In extremely difﬁ cult species, a last resort would be the micro-

grafting of the somatic embryo-derived shoots on suitable in vitro

germinated seedlings of the same species (and ideally also

genotype).

3 Two Example Protocols of Somatic Embryogenesis

As an example, optimized protocols for the induction of somatic

embryo genesis in Arabidopsis thaliana and Medicago truncatula

are reported here.

All stock solutions are prepared using ultrapure water (Milli-Q,

prepared by purifying deionized water to attain a sensitivity of 18

MΩ cm at 25 °C) and, unless stated otherwise, analytical grade

reagents. All reagents were purchased from Kalys (fr.kalys.com)

or Sigma-Aldrich (

www.sigmaaldrich.com ) and disposable plastic

ware (culture dishes, multiwall plates, pipettes, etc.) from

Dutscher (

www.dutscher.com ). When dealing with GMO mate-

rial, all needed precautions in terms of biosecurity are respected.

Reagents and stock solutions are generally stored in the fridge or

frozen until use, while media are kept at room temperature in the

dark until use.

For studies with Arabidopsis thaliana , seeds of wild-types C24 and

Col but also of the cytokinin-overproducing mutants hoc [

56 ] and

amp1 [

57 ] are used. These seeds are stored in a cold chamber (4

°C, in darkness) until used. For experiments with Medicago trun-

catula , R108-1 genotype seeds are generally used as the source of

explants.

Table

1 details the composition of basal media and stock solu-

tions used to prepare them. For Arabidopsis thaliana , MPic

medium is based on MS medium [

58 ] and contains 0.2 mg/L

picloram and benzylaminopurine (BAP) at 0.5 mg/L (for leaves)

[

29 ] or 1.0 mg/L (for roots). The sequence of SH-based media

used for Medicago truncatula is based on N6 major salt formula

[

61 ], SH microelements and vitamins [ 62 ], 0.38 mM FeEDTA,

0.55 mM myoinositol , and having the pH adjusted to 5.8 prior

to autoclaving. They differ in their sucrose content, mineral

strength, and hormonal composition. SH3 contains 4 mg/L

2,4-dichlorophenoxyacetic acid (2,4-D) plus 0.5 mg/L BAP and

30 g/L sucrose, while media SH9 and ½ SH9 are both hormone-

free and only differ in the salt strength which is reduced by half

3.1 Materials

3.1.1 Plant Materials

3.1.2 Composition

of Media Used

Sergio J. Ochatt and Maria Angeles Revilla

531

for ½ SH9. For preparation of semisolid media, SH3 medium is

further supplemented with 3 g/L Phytagel , while media SH9 and

½ SH9 are gelled with 7 g/L HP696 agar (Kalys).

A very straightforward protocol will permit the establishment of

the highly embryogenic culture in Arabidopsis , detailed in Fig.

1 ,

as follows:

1. Seeds of Arabidopsis wild-types C24 and Col but also the

cytokinin- overproducing mutants hoc [

56 ] and amp1 [ 57 ] are

sterilized during 10 min with a calcium hypochlorite solution

(2.5 %, w/v) and imbibed at 4 °C for 3 days before germina-

tion in constant light at 22 °C.

2. Germinated plants are placed in MS [

58 ] and grown in a

growth chamber at 22 °C with a 16/8 h day/night photope-

riod of a light intensity of 90 μmol/m

2 /s.

3. Plant age is calculated from the ﬁ rst day at 22 °C; 27 days after

germination , plants are harvested and individual leaves are

wounded with a scalpel, and roots cut to 1–2 cm length.

4. Leaf/root explants are kept stationary in liquid MS and MPic

media (pH 5.6), respectively, to regenerate embryos.

3.2 Methods

3.2.1 Arabidopsis

thaliana

Table 1

Composition of media and stock solutions

N6 Macro-salts (58)

(for 1 L of 10¥ stock soluon)

Medium SH9 (for 1 L); if

solidiﬁed add 7 g/L agar

Medium SH3 (for 1 L); if solidiﬁed

add 3 g/L Phytagel

Medium 0.5¥ SH9 (for 1 L);

if solidiﬁed add 7 g/L agar

Medium SHb10 (for 1 L);

if solidiﬁed add 6 g/L agar

Vitamins

(for 100 mL of 1000¥ soluon)

Stock soluons for SH Medium (59)

Micro-salts

(for 100 mL of 1000¥ soluon)

Macro-salts

(for 500 mL of 20¥ soluon)

From Stress to Embryos

532

5. Cultures are observed weekly during 6 weeks, and the medium

is not renovated until the end of the experiments, i.e., until

63 days.

6. By 6 weeks from culture initiation, cultures are transferred for

3 weeks onto hormone-free MS medium for expression of

embryogenesis.

7. After 9 weeks from culture initiation, cultures exhibit large

numbers of globular somatic embryo s and are transferred to

liquid medium of the same composition as used to induce

embryogenesis, cultured with shaking (80 rpm) under the

same conditions as above, and subcultured every 4 weeks

thereafter.

8. By the ﬁ rst passage with shaken conditions, the tissues from

both explant sources are completely covered with somatic

embryo s at different stages of development. The somatic

embryos start to detach from the explants, and 1 month later

the ﬂ asks will mostly contain germinating embryos only.

The barrel medic, Medicago truncatula , is considered as a model

legume species in terms of biotechnology approaches, and R108 is

one of the genotypes that can be regenerated by somatic embryo-

genesis and genetically transformed [

59 ], even if gene transfer is

efﬁ cient only for a few genotypes.

1. Use seeds of the R108-1 genotype of M. truncatula that have

been stratiﬁ ed (48–72 h in the dark at 4 °C), as described in

Trinh et al. [

60 ] and Ochatt et al. [ 16 ].

2. Scarify seeds (1 M H

2 SO

4 for 2 min or grated with sand paper)

for efﬁ cient germination .

3. Germinate seeds on humid ﬁ lter paper at 24 °C in the dark for

48–72 h.

4. Transfer germinated seeds to hormone-free MS medium [

16 ]

or SHb10 medium (Table

1 ; 60 ), at 22 ± 2 °C under a 16 h

light photoperiod at 90 μmol/m

2 /s from warm white ﬂ uores-

cent tubes, for 4–6 weeks.

5. Use individual folioles from trifoliate leaves (i.e., the cotyle-

donary leaves were discarded), harvested from the 4–6-week-

old plantlets grown in vitro ( see step 4 ). Alternatively, folioles

from trifoliate leaves on in vivo seedlings of the same age, but

grown in the glasshouse (19–22 °C, 60–70 % relative humid-

ity, 16/8 h light/dark photoperiod at 200 μmol/m

2 /s) may

also be used.

6. For the initiation of callus followed by somatic embryo induc-

tion, folioles are cut 3–4 times with the blade of a scalpel (per-

pendicularly to the main vein) and transferred to SH3 medium,

which contains 3 % (w/v) sucrose (commercial sugar can be

3.2.2 Medicago

truncatula (Table 1 )

Sergio J. Ochatt and Maria Angeles Revilla

533

used) and is also supplemented with 16 μM (4 mg/L) 2,4-D

and 2 μM (0.5 mg/L) BAP. Cultures are kept at 25 °C in the

dark, and explants transferred to fresh medium every 2 weeks.

7. During 8–10 weeks of culture, callus develops and some early

globular pro-embryonic structures appear on the callus

surface.

8. After 8–10 weeks, callusing explants are transferred to

hormone- free medium SH9, which contains only 2 % (w/v)

sucrose , and cultures are transferred to the photoperiodic light

and temperature regime as reported above for expression of

somatic embryo genesis. On this medium, somatic embryos

develop up to the cotyledonary stage and start to germinate.

9. Transfer germinating somatic embryo s to half-strength SH9

medium for complete rooting of plantlets, which are thereafter

acclimatized and transferred to the glasshouse until maturity

and seed set.

It is important to note that if this protocol is used for gene

transfer or in vitro selection for stress tolerance, once a given callus

produces a shoot from embryos, this is harvested and the callus is

discarded to prevent multiplication of several plants derived from

the same transformation or selection event.

4 Conclusions

The FAO [ 63 ] has been ringing the alert about the ever-increasing

demand for food and feed resulting from the constant demo-

graphic increase, while most quality arable land is already under

exploitation, which pushes cultivation to more marginal areas and

soils so that crops are confronted with novel or increased stress

agents [

64 ]. The need for developing new more stress-resistant

genotypes to ensure food supply is becoming urgent, and one of

the ways to producing them is by exploiting biotechnology

approaches in vitro, such as somatic embryo genesis. Against this

background, integration of conventional breeding programs and

molecular and cell biology approaches based on somatic embryo-

genesis proves invaluable to fasten the generation of genetically

improved commercial crop species [

10 , 13 , 49 , 64 ].

Despite all the knowledge about the requirements for in vitro

regeneration accumulated in the literature over the last decades, it

is still a matter of controversy why certain genotypes, cells, or

explants are embryogenic, while others are not [

13 , 15 , 19 ]. Thus,

the optimization of culture conditions remains mainly an empirical

exercise driven by experience and intuition of researchers to assess

a range of combinations of potentially effective parameters.

However, various recent advances would permit to better deﬁ ne

From Stress to Embryos

534

and measure the totipotent status and hence the degree of cell

speciﬁ city toward regeneration competence. In this context, it is

most likely that the reason for this gap in knowledge comes from

the complex interactions in place between the environmental con-

ditions (including the composition of the used culture medium)

and the physiological status of the cells at the time of culture which

are under the tight control of genes whose expression also depends

on many factors and conditions [

6 , 13 , 19 , 45 ].

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Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

Chapter 32

Cryotechniques for the Long-Term Conservation

of Embryogenic Cultures from Woody Plants

Elif Aylin Ozudogru and Maurizio Lambardi

Abstract

Since its development in the 1960s, plant cryopreservation is considered an extraordinary method of safe

long-term conservation of biological material, as it does not induce genetic alterations and preserve the

regeneration potential of the stored material. It is based on the storage of explants at cryogenic temperatures,

such as the one of liquid nitrogen (−196 °C), where the metabolism within the cells is suspended; thus, the

time for these cells is theoretically “stopped”. Cryopreservation is particularly important for embryogenic

cultures, as they require periodic subculturing for their maintenance, and this, in turn, increases the risk of

losing the material, as well as its embryogenic potential. Periodic re-initiation of embryogenic cultures is pos-

sible; however, it is labor intensive, expensive, and particularly difﬁ cult when working with species for which

embryogenic explants are available only during a limited period of the year. Among various methods of

cryopreservation available for embryogenic cultures, slow cooling is still the most common approach, espe-

cially in callus cultures from softwood species. This chapter brieﬂ y reviews the cryopreservation of embryo-

genic cultures in conifers and broadleaf trees, and describes as well a complete protocol of embryogenic callus

cryopreservation from common ash tree ( Fraxinus excelsior L.) by slow cooling.

Key words Cryopreservation , Embryogenic cultures , Fraxinus excelsior L. , Slow cooling , Somatic

embryo s , Two-step freezing , Woody trees

1 Introduction

Cryopreservation refers to the storage of biological material (such

as seeds, somatic/ zygotic embryo s, embryogenic and organogenic

callus cultures, shoot tips, axillary buds) in liquid gases, at ultra-

low temperatures. Among various gases available, liquid nitrogen

( LN ), which is in such a physical state at −196 °C, is usually pre-

ferred due to its numerous advantages, such as low cost, ease in

handling, ease in delivery, and no toxicity for preserved plant speci-

mens. When a biological material is subjected to such a low tem-

perature, almost all of the biological reactions within its cells are

hampered. Suspension of the metabolism in such way ensures the

storage of the material for theoretically unlimited time, without

inducing any genetic alteration. Hence, the technique is considered

538

the only valuable approach to the ex situ long-term conservation of

plant biodiversity that can be regarded as complementary to the

traditional storage in seed banks and in in-ﬁ eld collections [

1 ].

Water content of the cells during the immersion in LN is the key

factor affecting the success of cryopreservation. It should be low

enough to prevent the formation of lethal intracellular ice crystals

and yet high enough to enable recovery of viable explants after

storage in LN. This is induced by triggering the “vitriﬁ cation” of

cytosol, which is a physical state of water solutions, in which the

solidiﬁ cation of water molecules at ultra-low temperature is

obtained through their transition to an amorphous (“glassy”)

state, instead of crystallization [

2 ]. Cell vitriﬁ cation can be induced

in several ways: (1) by imposing “ cryodehydration ” of explants,

i.e., by a gradual decrease of their temperature (usually at a rate of

−0.5/−1 °C/min) up to −40 °C before the immersion in LN

(“ slow cooling ”), (2) by the use of highly concentrated vitriﬁ ca-

tion solutions (“ chemical dehydration ”), or (3) by exposing the

explants to sterile air ﬂ ow or silica gel (“ physical dehydration ”).

The latter two techniques allow the direct immersion of the

explants in LN and thus are referred as “ one-step freezing ” [

3 ].

Cryopreservation is particularly important for embryogenic

cultures because, once a cell culture is established, it requires peri-

odic subculturing for the maintenance, and this is not only labor

intensive but also increases the risk of losing either the material

through contamination, human errors or technical failures, or its

embryogenic potential through the frequent long-term subcultur-

ing [

4 , 5 ]. Periodic re-initiation of embryogenic cultures can pro-

vide a solution to this drawback. However, this is labor intensive

and expensive, too, and is particularly difﬁ cult when working with

species for which suitable explants for embryogenic callus induc-

tion are available only during a limited period of the year [

6 – 8 ].

Nevertheless, development and optimization of efﬁ cient cryo-

preservation protocols for embryogenic cultures allow the safe,

low-cost, and long-term conservation of this unique material [

9 ].

Cryopreservation of embryogenic cultures is a relatively recent

application of the cryogenic technology, ﬁ rst examples of detailed

and successful protocols being available only in the early 1990s

[

10 , 11 ]. Sakai’s work on Citrus sinensis also reported the develop-

ment of Plant Vitriﬁ cation Solution no. 2 ( PVS2 ), a mixture of

cryoprotectants which instantly became a milestone to induce cell

vitriﬁ cation . It was soon evident that the possibility of storing valu-

able embryogenic culture lines in LN could allow the long-term

maintenance of their embryogenic potential, making them avail-

able only when necessary and avoiding the above-mentioned draw-

backs induced by repeated subculturing [

9 ].

Slow cooling is the most common approach for embryogenic

callus cultures. In recent years, this approach allowed the develop-

ment of effective protocols for various conifer (Table

1 ) and broad-

leaf (Table

2 ) trees. One disadvantage of slow cooling is the

Elif Aylin Ozudogru and Maurizio Lambardi

Table 1

Cryopreservation of embryogenic cultures of conifers ( LN liquid nitrogen , NR not reported, RT room temperature)

Species Explant a Preculture Pre-freezing treatments Cooling rate Thawing

Maximum

recovery (%) Reference

Slow cooling

Abies cephalonica CC 5 °C (14 days)+,

darkness + sucrose, 0.2 M

(1 day) + 0.4 M (1 day)

10 % PEG6000+ 10 %

glucose + 10 % DMSO

30 min, 0 °C

−10 °C/h to −38 °C 37 °C 75 [

16 ]

Abies hybrids CC Sorbitol, 0.4 or 0.8 M

(2 or 3 days) 0.4 or 0.8 M sorbitol +

5 % DMSO, 1 h, 0 °C −80 °C (~100 min)

to −40 °Cc

40 °C, 3 min 37–100 [ 17 ]

Picea abies SC Sorbitol, 0.4 M (2 days) 5 % DMSO, 30 min −0.3 °C/min to

−35 °C 37 °C, 2–3 min NR [

18 ]

Picea abies SC Sorbitol, 0.2 M (1 day) +

0.4 M (1 day), darkness 5 % DMSO, 0 °C −0.5 °C/min to

−40 °C 45 °C, 90 s NR [

19 ]

Picea abies CC Sorbitol, 0.4 M (2 days) 5 % DMSO, 30 min −0.3 °C/min to

−35 °C 37 °C, 2–3 min NR [

20 ]

Picea glauca

engelmannii SC – 0.4 M sorbitol + 5 % DMSO Multistep cooling

b 37 °C, 1–2 s 100 [ 21 ]

Picea glauca SC Sorbitol, 0.4 M (1 day) 0.4 M sorbitol + 5 % DMSO,

30 min −0.3 °C/min to

−35 °C 37 °C, 90 s 94 [

22 ]

Picea sitchensis SC Sorbitol, 0.2 M (1 day) +

0.4 M (1 day) 5 % DMSO, 0 °C −0.5 °C/min to

−40 °C 45 °C NR [

23 ]

Picea sitchensis SC Sorbitol, 0.2 M (1 day) +

0.4 M (1 day), darkness 5 % DMSO, 0 °C −0.5 °C/min to

−40 °C 45 °C, 90 s NR [

19 ]

Pinus caribaea

‘Hondurensis’ SC Sucrose, 0.4 M 0.4 M sucrose + 5 % DMSO −0.5 °C/min to

−35 °C 40 °C, 2 min 100 [

24 ]

Pinus nigra CC Sucrose or maltose, 0.5 M

(1 day) 0.5 M sucrose or maltose +

7.5 % DMSO, 1 h −80 °C (~100 min)

to -40 °C 40 °C, 3–4 min 87 [

25 ]

Pinus patula SC Sorbitol, 0.3 M (1 day) 0.3 M sorbitol + 5 % DMSO,

20 min, 0 °C −70 °C, 2 h

c 42 °C, 2–3 min 60 [ 26 ]

Pinus pinaster SC Maltose, 0.2 M (1 day) +

0.4 M (1 day), darkness 10 % PEG4000 + 10 %

sucrose + 10 % DMSO 0 °C −80 °C, 1 day

c 45 °C 97 [ 27 ]

(continued)

Table 1

(continued)

Species Explant a Preculture Pre-freezing treatments Cooling rate Thawing

Maximum

recovery (%) Reference

Pinus pinaster SC Sucrose, 0.22 M (1 day) +

0.4 M (2 h) 5 % PSD solution (20 g/L

PEG4000 + 20 g/L

sucrose + 20 % (v/v)

DMSO) , 1 h at 0 °C

−70 °C, 80 min

c 40 °C 100 [ 28 ]

Pinus radiata SC Sorbitol, 0.4 M (1 day) 20 °C 0.4 M sorbitol +10 %

DMSO −80 °C, 75–90 min

c RT, 30 s + 40–45

°C, 2 min 100 [ 29 ]

Pinus roxburghii SC Sorbitol, 0.3 M (1 day) 0.3 M sorbitol +5 % DMSO Multistepcooling

d 45 °C, 2–3 min 70 [ 30 ]

Pinus sylvestris CC 5 °C (14 days) + sucrose,

0.2 M (1 day) + 0.4 M (1 day) 10 % PEG6000+ 10 %

glucose + 10 % DMSO, 30

min, 0 °C

−10 °C/h to −38 °C 37 °C 78 [

31 ]

Pinus sylvestris CC Sucrose, 0.2 M (1 day) +

0.4 M (1 day) 10 % PEG6000 + 10 %

glucose + 10 % DMSO,

1 h, 0 °C

−10 °C/h to −38 °C 37 °C, 1–3 min 80–93 [

32 ]

Direct immersion in LN

Picea mariana CC Sorbitol, 0.8 M (2 days) PVS2, 30 min, 0 °C – 40 °C 67 [

33 ]

Updated from [ 9 ]

a CC clumps of embryogenic callus (developmental stage not reported), SC cells from suspension cultures

b Slow cooling was achieved as follows: −0.3 °C/min to −3 °C → −15 °C/min to −8 °C → −25 °C/min to −32 °C → −0.3 °C/min to −35 °C

c Slow cooling was achieved by incubating the samples in a freezer at −70 or −80 °C before plunging them into LN

d Slow cooling was achieved as follows: −0.3 °C/min to −35 °C → −25 °C/min to −50 °C

Table 2

Cryopreservation of embryogenic cultures of broadleaf trees ( LF laminar ﬂ ow, LN liquid nitrogen , MC moisture content, NR not reported, RT room temperature, SG

silica gel)

Species Explant a Preculture Pre-freezing treatments Cooling rate Thawing

Maximum

recovery (%) Reference

Slow cooling

Citrus deliciosa SC – 10 % DMSO, 30 min, 4 °C −0.5 °C/min 37 °C, 5 min NR [ 34 ]

Citrus sinensis SE – – −0.5 °C/min to −42 °C RT, 15 min 5 [ 35 ]

Citrus sinensis HSE – 10 % DMSO −0.5 °C/min to −42 °C 37 °C, 5 min 31 [ 36 ]

Citrus ssp. CC – 10 % DMSO, 30 min, 0 °C −0.5 °C/min to −40 °C 37 °C, 5 min 100 [ 37 ]

Fraxinus excelsior CC – 0.61 M sucrose +7.5 %

DMSO, 1 h, 0 °C −1 °C/min to −40 °C 40 °C, 2-3 min 100 [ 12 ]

Hevea brasiliensis CC – 1 M sucrose + 10 % DMSO,

1 h, 0 °C −0.2 °C/min to −40 °C 40 °C 49 [

38 ]

Hevea brasiliensis CC – 1 M sucrose + 10 % DMSO,

1 h, 0 °C −0.2 °C/min to −40 °C 40 °C 70 [

39 ]

Persea americana CC – 5 % DMSO + 5 %

glycerol + 0.13 or 1 M

sucrose, 30 min, 0 °C

−1 °C/min to −80 °C 40 °C, 5 min NR [

40 ]

(continued)

Species Explant a Preculture Pre-freezing treatments Thawing

Maximum

recovery (%) Reference

Direct immersion in LN

Aesculus

hippocastanum TSE 4 °C (5 days), darkness 2 M glycerol + 0.4 M sucrose, 30 min,

25 °C+ PVS2, 90 min, 0 °C 45 °C, 50 s 94 [

41 ]

Citrus sinensis SC – 60 % PVS2, 5 min, 25 °C+ PVS2,

3 min, 0 °C 25 °C 84 [

11 ]

Citrus ssp. SE Sucrose, 0.7 M (1 day) Encapsulation + dehydration, 5 h, LF RT, 2–3 min 100 [

42 ]

Fraxinus

angustifolia CSE 4 °C (10 h) → sucrose,

0.5 M (1 day) Encapsulation + dehydration, 1 h, SG 40 °C, 3 min 31 [

43 ]

Litchi chinensis CC – PVS2, 30 min, 0 °C 38–40 °C, 2 min 100 [

44 ]

Mangifera indica

‘Zihua’ SC Sucrose, 0.5 M (1 day)

25 °C PVS3

b , 20 min, 25 °C 25 °C, 2–3 min 94 [ 45 ]

Olea europaea

‘Canino’ CC 4 °C (4 days), darkness 2 M glycerol + 0.4 M sucrose, 30 min,

25 °C+ PVS2, 90 min, 0 °C RT, 10 s + 40 °C, 50 s 38 [

46 ]

Olea europea

‘Nabali’ CC 30 °C (1 day) Encapsulation + 0.4 M sucrose +

2 M glycerol, 60 min + PVS2, 3 h, 0 °C 38 °C, 2 min 64 [

47 ]

Olea europea

‘Picual’ CC Sucrose, 0.4 M

(7–8 weeks) PVS2, 60 min, 0 °C, droplet vitriﬁ cation 1.2 M sucrose, RT 100 [

48 ]

Persea americana CC – PVS2, 60 min 40 °C, 5 min NR [

40 ]

Prunus avium CC Multistep sucrose

preculture

c Dehydration, LF (until 20 % MC) 1.2 M sucrose at 40 °C 89 [

49 ]

Quercus robur GSE- HSE Sucrose, 0.3 M (3 days) PVS2, 60 min, 25 °C 40 °C, 2 min 70 [

50 ]

Quercus robur CC Multistep sucrose

preculture

c Dehydration, 4–5 h, LF 1.2 M sucrose at 40 °C NR [

51 ]

Quercus suber GSE Sucrose, 0.3 M (3 days) PVS2, 60 min, 0 °C 40 °C, 2 min 93 [

52 ]

Quercus suber GSE Sucrose, 0.7 M (3 days) Dehydration, LF (until 25–35 % MC) 38 °C, 2 min 90 [

53 ]

Theobroma cacao SSE Sucrose, 0.5 M (5 days) PVS2, 60 min, 0 °C 42 °C, 3 min 74.5 [

54 ]

Updated from [ 9 ]

a CC clumps of embryogenic callus (developmental stage not reported), CSE clumps or isolated somatic embryo s at the cotyledonary stage, GSE clumps or isolated somatic embryos at the

globular stage, HSE clumps or isolated somatic embryos at the heart stage, SC cells from suspension cultures, SSE secondary somatic embryos, SE isolated somatic embryos, TSE clumps or

isolated somatic embryos at the torpedo stage

b PVS3: 50 % sucrose (w/v) + 50 % glycerol (w/v) in standard culture medium

c Multistep sucrose preculture was performed as follows: sucrose, 0.25 M (1 day) + 0.5 M (1 day) + 0.75 M (2 days) + 1.0 M (3 days)

Table 2

(continued)

543

requirement of an expensive equipment, the controlled-rate

freezer. The Nalgene freezing container “Mr. Frosty”

® (Sigma-

Aldrich) is a cheaper alternative approach to slow cooling; how-

ever, it should be noted that “Mr. Frosty”

® provides only a rate of

−1 °C/min gradual temperature decrease and thus is useful only

when this cooling rate is suitable for the plant material [

9 ].

Vitriﬁ cation- based protocols have also been developed for embryo-

genic cultures from various important plant species, such as Citrus

spp., Olea europaea , Fraxinus spp., and Quercus spp. (Tables

1 and

2 ). A sample protocol on cryopreservation of Fraxinus excelsior

embryogenic callus cultures by slow cooling approach is included

to the chapter, providing a detailed information of the procedure

to the reader.

2 Materials

Slow cooling of Fraxinus excelsior is presented as a sample proto-

col [

12 ], where embryogenic callus cultures at the proliferation

phase (Fig.

1a ) are used as a plant material ( see Notes 1 and 2 ).

For embryogenic callus induction, following seed collection and

their decontamination, zygotic embryo s are isolated, and embry-

onic axes are excised aseptically and cultured on ½-strength MS

[

13 ] medium supplemented with 8.8 μM 2,4-dichlorophenoxy-

acetic acid (2,4-D) and 4.4 μM benzyladenine (BA) ( see Note 3 )

for 2 months in the dark, followed by an additional 1-month incu-

bation period on plant growth regulator ( PGR )-free medium

(MS0) under standard culture conditions, i.e., 23 ± 1 °C under a

16-h photoperiod and low light intensity (20 μmol/m

2 /s). The

de novo formed embryogenic callus is then transferred to Woody

Plant Medium (WPM,

[14] ) ( see Note 4 ), supplemented with 4.4

μM BA, and maintained by subculturing at 2-week intervals (see

also

[15] ).

1. Sucrose -rich liquid medium: Liquid WPM, containing 0.61 M

(210 g/L) sucrose ( see Note 5 ).

2. Dimethyl sulfoxide ( DMSO ) and sucrose-rich liquid medium:

Liquid WPM, containing 0.62 M (210 g/L) sucrose and 15 %

DMSO (w/v) ( see Note 6 ).

3. Post-thaw recovery and plantlet development medium: Semi-

solid WPM supplemented with 4.4 μM BA.

4. Somatic embryo maturation medium: PGR -free semi-solid

WPM .

1. Graduate cylinders (100, 500, 1000 mL).

2. Glass beakers (250, 500, 1000 mL).

2.1 Plant Material

2.2 Cryoprotective

Solutions

and Semi- solid Media

2.3 Laboratory

Facilities

Cryostorage of Embryogenic Cultures

544

3. Magnetic stirrer and spin bar.

4. Analytical balance, weighting containers, and lab spoons.

5. pH meter.

6. Autoclave.

7. Horizontal ﬂ ow cabinet.

8. Automatic pipettor.

9. Nalgene benchtop cooler.

10. Nalgene freezing container (“Mr. Frosty”

® , Sigma-Aldrich).

11. Nalgene box.

12. LN dewar.

13. Appropriate gloves and masks for protection from LN .

14. Growth chamber (temperature of 23 ± 1 °C, photosynthetic

photon ﬂ ux density of 20 μmol/m

2 /s and 16-h photoperiod).

1. Distilled water.

2. LN .

3. NaOH and HCl solution (1.0 and 0.1 N).

2.4 Consumables

Fig. 1 Slow cooling of embryogenic callus cultures of Fraxinus excelsior . ( a ) Embryogenic callus cultures at the

proliferation phase used in the cryopreservation trials. ( b ) Callus samples (~2 g) transferred to sterile 10-mL

glass tubes for the treatment with cryoprotective solutions. ( c ) Distribution of the mixture into sterile 2-mL

cryovials. ( d ) “Mr. Frosty”

® containing cryovials to be cooled in a −80 °C freezer. ( e ) Immersion of the samples

in LN . ( f ) Cryopreserved callus samples immediately after thawing and placing on post-thaw recovery medium.

( g ) Proliferation of the callus samples 42 days after thawing and recovery. ( h ) Somatic embryo maturation on the

cryopreserved callus samples ( arrows ) (Figg. g and h reproduced from [ 12 ] with permission from CryoLetters)

Elif Aylin Ozudogru and Maurizio Lambardi

545

4. Stock solutions of macro- and microelements, organics, and

iron of MS and WPM media .

5. Stock solutions of growth regulators (2,4-D and BA).

6. Sucrose .

7. Gelrite .

8. Schott bottles (250, 500, 1000 mL).

9. Petri dishes (Ø 90 mm).

10. 50-mm Whatman ﬁ lter paper.

11. Forceps, scalpels, and blades.

12. Sterile medical gloves.

13. Sterile tubes (10 mL).

14. Sterile cryovials (2 mL).

15. Pipettes.

16. Trays (35 mm Ø).

3 Methods

Cryopreservation of embryogenic callus cultures (as it is for cryo-

preservation of almost all kinds of plant material) is a multistep

process. It involves several consecutive preconditioning (to enhance

the cold tolerance of plant material before the immersion in LN )

and promotive steps (to help the plant material recovering after

storage). Although some of these steps can be skipped, depending

on the cryopreservation approach applied, a complete cryopreser-

vation protocol is composed of (1) cold hardening, (2) preculture,

(3) osmoprotection, (4) cryoprotection , (5) immersion and stor-

age in LN, (6) thawing, (7) rinsing, and (8) plating on regenera-

tion medium.

Cryopreservation approach proved to be suitable for the

embryogenic callus cultures of Fraxinus excelsior is based on slow

cooling , inducing “ cryodehydration ” ( see Note 7 ) of the samples.

Here, the main step is the gradual decrease of the temperature at a

rate of −0.5/−1 °C/min to an intermediate temperature of −40 °C

before the immersion in LN , while some of the steps preceeding

immersion in LN (i.e., cold hardening, preculture, and osmopro-

tection) can be skipped. Gradual decrease of the temperature of

the samples, inducing cryodehydration, can be achieved by using a

controlled-rate freezer or the Nalgene freezing container “Mr

Frosty”

® , a specially designed plastic box containing 250 mL iso-

propyl alcohol, which cools the samples at a rate of about −1 °C/

min [

3 , 9 ].

Cryostorage of Embryogenic Cultures

546

1. Embryogenic callus samples (~2 g) are placed in sterile 10-mL

glass tubes (Fig.

1b ) and incubated with 5 mL sucrose -rich

liquid medium, i.e., liquid WPM, containing 210 g/L (0.61

M) sucrose.

2. Five mL DMSO and sucrose -rich liquid medium, added of 15

% DMSO, is then gradually added in three steps (1 mL, 2 mL,

and 2 mL, respectively) over a total period of 60 min (15 min,

15 min, and 30 min, respectively), to reach a ﬁ nal DMSO con-

centration of 7.5 % in a ﬁ nal volume of 10 mL.

3. Following DMSO treatment, suspension cultures are mixed

thoroughly and transferred into sterile 2-mL cryovials (each

cryovial containing 1 mL of the mixture, Fig.

1c ) ( see Note 8 ).

4. Slow cooling of the samples is achieved by placing the cryovials

in the cells of “Mr. Frosty”

® (Fig. 1d , see Note 9 ) and transfer-

ring the device in a −80 °C freezer, where it is kept (for about

1 h) until the temperature reaches −40 °C.

5. Afterward, the cryovials are rapidly transferred to Nalgene

boxes that are then plunged into LN (Fig.

1e ) where they are

stored for at least 1 h.

6. Callus samples that are treated with DMSO (control 1), or

treated with DMSO and cooled to −40 °C, but not frozen in

LN (control 2) serve as control.

1. The embryogenic callus samples are thawed in a 40 °C water

bath until the DMSO solution is totally melted.

2. They are then poured onto a 50-mm Whatman ﬁ lter paper,

placed on post-thaw recovery medium, i.e., WPM supple-

mented with 4.4 μM BA (Fig.

1f ), and cultured at 23 ± 1 °C in

the dark for 2 days.

3. Subsequently, embryogenic callus samples are subcultured

under standard culture conditions (i.e., 23 ± 1 °C, 16-h photo-

period, with a light intensity of 20 μmol/m

2 /s) in every

2 weeks by transferring the ﬁ lter paper onto semi-solid fresh

post-thaw recovery medium until the 42nd day (Fig.

1g ).

4. The callus clumps are then put in direct contact with fresh

semi- solid medium.

1. In order to stimulate somatic embryo maturation and conver-

sion into plantlets, cryopreserved callus samples are transferred

onto PGR -free, semi-solid WPM ( embryo maturation medium)

and subcultured at 2-week intervals (Fig.

1h ).

2. Somatic embryo s at the cotyledonary stage are then isolated

and subcultured on the same medium at 4 °C, in darkness for

4 weeks, followed by transfer to WPM containing 4.4 μM BA

under the above-mentioned standard culture conditions.

3. The plantlets developed are then transferred to trays (35 mm

Ø) and acclimatized under greenhouse conditions.

3.1 Cryopreservation

of Fraxinus excelsior

3.2 Thawing

and Post-Thaw

Recovery

3.3 Embryo

Maturation

Elif Aylin Ozudogru and Maurizio Lambardi

547

4 Notes

1. For cryopreservation trials, only established embryogenic cal-

lus cultures (i.e., coming from cultures maintained for at least

1 year) are used.

2. Cultures were initiated from immature zygotic embryo s of

selected ash trees in Florence, Italy [

15 ].

3. In our laboratory, stock solutions of PGR are prepared in 10

−3

M concentration and are stored at 4 °C. 2,4-D and BA are

sterilized by autoclaving; thus, they are included in the medium

before sterilization.

4. MS and WPM media , supplemented or not with PGR , con-

tained 20 g/L sucrose and 3.6 g/L Gelrite (pH 5.8).

5. During the course of the study, sucrose concentration, as well

as the application time of the solution, should be optimized

carefully for each species. Accordingly, depending on the plant

species used in the study, the solution can be prepared using

different basal medium formulations.

6. As stated in Note 5 , also here sucrose concentration, as well as

DMSO concentration, should be optimized carefully accord-

ing to the sensitivity of the speciﬁ c embryogenic callus line.

The solution can be prepared in any kind of basal medium

formulation. However, what is crucial is that the sucrose con-

centrations and basal medium formulations of these two solu-

tions should be identical.

7. “ Cryodehydration ” refers to a state of losing moderate amount

of potentially freezing water molecules from the cell cytosol, in

response to gradual decrease of the temperature. If tempera-

ture decrease is performed too fast, cells do not lose sufﬁ cient

amount of water, and thus they risk the cryo-damages induced

by ice crystals formed in the intra cellular spaces during immer-

sion in LN . On the contrary, if temperature decrease is per-

formed too slowly, cells lose extreme amount of water, which

results in dehydration injuries due to cell plasmolysis. Moderate

water loss is proved to be induced by decreasing the tempera-

ture at a rate of −0.5/−1 °C/min.

8. This application is rather “tricky” while working with compact

callus samples, as it is difﬁ cult to have homogeneous distribu-

tion of callus samples in the solution and thus is difﬁ cult to

transfer equal amount of callus sample from the 10-mL glass

tubes into each 2-mL cryovial. Alternatively, callus samples of

equal amount can be directly transferred to cryovials and can

be treated with the cryoprotective solutions in that container.

If this is the case, it should be recalled that the ﬁ nal volume of

the solutions will be 1 mL, in total.

Cryostorage of Embryogenic Cultures

548

9. “Mr. Frosty”

® has a plate of 18 cells. Thus, it should be consid-

ered that, using one “Mr. Frosty”

® container, only 18 cryovials

can be managed for each trial.

Acknowledgment

The “Ente Cassa di Risparmio of Florence” is gratefully acknowl-

edged for a 36-month grant to Elif Aylin Ozudogru (project

“POLICENTRO”).

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Elif Aylin Ozudogru and Maurizio Lambardi

551

A

Abies alba ...........................................70, 72, 89, 90, 417, 429

Abies cephalonica (methods)

cryopreservation by controlled-rate cooling ................426

somatic embryogenesis ...............................................424

Abscissic acid (ABA) .............................. 11, 29, 32, 33, 35, 36,

38, 39, 41, 42, 58, 62, 68, 70, 74, 75, 132, 136–138,

141, 142, 144–146, 150, 155, 158, 173, 177, 178,

185, 189, 195, 198–200, 215, 408, 409, 422–425,

443, 445, 449, 524

Acca sellowiana ....................................................34, 122, 125

Acrylic resin sections ........................................................493

Activated charcoal (AC) ........................... 5, 74, 94, 119, 133,

135, 136, 138, 145, 146, 156, 158, 177, 218, 230,

267, 268, 281, 282, 311, 313, 332, 348, 380, 381,

408, 409, 412, 432, 434, 435, 437, 443, 445, 525

Adventitious embryony ........................................... 4, 5, 8, 9,

15, 16, 294

Aesculus hippocastanum (methods)

acclimatization ............................................................437

culture media and conditions ......................................434

explant surface sterilization .........................................435

somatic embryo development and embryogenic tissue

proliferation ....................................................435

somatic embryo maturation and conversion ...............435

somatic embryogenesis induction ...............................435

Agar .......................................... 158, 210, 214, 230, 266–268,

273, 281, 293, 296, 309–313, 316, 317, 332, 334,

343–345, 347, 361, 363, 373, 375, 388, 389, 391,

393, 397, 399, 422, 426, 432, 435, 454, 463, 464,

470, 478, 479, 485, 518, 519, 529, 531, 545

Agrobacterium .............................. 25, 265, 267, 268, 271–273,

303, 305, 306, 309, 316–317

Agrobacterium mediated transformation ............ 303, 305, 306

Air-lift ...................................................................... 158, 249

Albino plants ....................................................................400

Alternative oxidase enzyme (AOX) ........................ 90, 93–95

Anderson medium ............................................................332

Androgenesis .......................... 7, 69, 104, 209, 212, 217, 219,

223, 228, 231, 232, 454, 467

Angiosperms .............................................6, 9, 26, 53, 68, 70,

72, 75, 89, 103, 104, 122, 167, 169, 175, 180, 185,

198, 525

Anther culture ..........................210, 212, 214, 217–220, 222,

225, 226, 229, 230, 234, 296, 453, 454, 456, 457,

460, 464, 467, 468, 470–473, 476, 478, 480–482,

484, 485, 493, 495, 496

Anther filament .................................432, 434, 436, 463, 473

Antigenicity ......................................................................492

Apomixis ............4, 5, 6, 8, 9, 12, 13, 15, 17, 19, 63, 294, 395

Arabidopsis thaliana .......................... 26, 47, 74, 167, 530–532

Arabinogalactan proteins (AGPs) ............ 41, 91, 124, 178, 227

Araucaria angustifolia (methods)

early somatic embryos maturation ..............................447

embryogenic callus pre-maturation 1..........................447

embryogenic callus pre-maturation 2..........................447

embryogenic callus proliferation .................................446

morphological and cytochemical analysis

procedure ........................................................448

somatic embryogenesis induction ...............................446

Araucariaceae ..................................................... 131, 146, 439

Arbutus unedo ............................................................ 329, 337

ARGONAUTE (AGO) genes ..................................53, 183

Artificial endosperm ......................................... 516–519, 521

Artificial seed See Synthetic seed

Aascorbic acid............................307, 338, 478, 506, 517, 518

Asexual seed .........................................................................4

Auxins

2,4-dichlorophenoxi acetic acid

(2,4-D)...................................51, 58, 69, 524, 530

3-indolbutyric acid (IBA) ...........................................530

1-naphthaleneacetic acid (NAA) ................................530

B

BABY BOOM (BBM) gene .................................................14

Bactris gasipaes (methods)

conversion of somatic embryos and plantlet

acclimatization ................................................286

somatic embryogenesis from inflorescences ................283

somatic embryogenesis from zygotic

embryos ..........................................................282

use of thin cell layers as explants to induce somatic

embryogenesis .................................................285

Ballon column reactor (BCR) .........................245, 246, 249,

251, 252

Balloon-type bubble reactor (BTBR) ............ 245, 248–252, 255

INDEX

Maria Antonietta Germanà and Maurizio Lambardi (eds.), In Vitro Embryogenesis in Higher Plants, Methods in Molecular Biology,

552

IN VITRO EMBRYOGENESIS IN HIGHER PLANTS

Index

Barley ..........................74, 108, 109, 211, 213, 468, 495, 503,

504, 507, 508, 510–511, see also Hordeum vulgare

Bax-inhibitor 1 (BI-1) .............................................. 109, 110

Beta-naphthoxyacetic acid (NOA) ................................... 267

Betula pendula ...................................................................123

Bioreactor ................................158, –159, 248–250, 252, 253

Bipolar propagules ............................................................ 516

Brachiaria brizantha ..................................................395–401

Brachiaria ruziziensis .........................................................400

Brassica napus ............................. 13, 32, 54, 62, 215, 218, 468

Bubble reactor ..........................................................249, 252

Bulblets ............................................................. 389, 391–393

Bulbs .................................. 306, 351, 352, 387, 389–391, 393

C

Calcium alginate matrix ...................................................520

Calcium chloride ...................................... 308, 517, 518, 520

Calcium ions .................................................................68, 70

Canola ...................................................................... 211, 292

Capsicum ................................................... 211, 228, 229, 468

Capsicum annum (methods)

analysis of the ploidy level ..........................................472

donor plant growth conditions ...................................469

in vitro culture of anthers ............................................470

Carbenicillin ..................................................... 268, 272, 274

Carbohydrates .............. 31–34, 38, 119, 124, 177, 192, 196, 378

Carbon dioxide (CO2) .......................158, 230, 247, 248, 413

Carrot ............................... 13, 17, 25, 29, 41, 51, 53, 58, 62, 63,

67–70, 89, 91, 93, 94, 118, 122, 123, 249, 252, 292,

In Vitro Embryogenesis in Higher Plants (1)

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