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Bruno Alexandre Valente da Silva Costa
EXTRACTION AND FRACTIONATION OF
CROWBERRY AND TAMARILLO LEAVES FOR
BIOLOGICAL ACTIVITY SCREENING
Dissertation submitted to the University of Coimbra for the fulfillment of
the necessary requirements for the obtention of the degree of Master in
Biodiversity and Vegetal Biotechnology, with scientific supervision of Dr.
Jorge Manuel Pataca Leal Canhoto (CEF, DCV) and Dr. Ricardo Manuel
Fernandes da Costa (MF-Q, CEF)
October 2021
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Extraction and fractionation of crowberry and
tamarillo leaves for biological activity screening
Extração e fracionamento de folhas de camarinha
e tamarilho para screening de atividade biológica
Bruno Alexandre Valente da Silva Costa
Department of Life Sciences of the Faculty of Sciences and Technology
Departamento de Ciências da Vida da Faculdade de Ciências e Tecnologia
Dissertation submitted to the University of Coimbra for the fulfillment of the necessary
requirements for the obtention of the degree of Master in Biodiversity and Biotechnology. Study
carried with scientific supervision of Dr. Ricardo Manuel Fernandes da Costa (MF-Q, CFE)
and Dr. Jorge Manuel Pataca Leal Canhoto (CEF, DCV).
Dissertação apresentada à Universidade de Coimbra para cumprimento dos requisitos
necessários à obtenção de grau de Mestre em Biodiversidade e Biotecnologia Vegetal. Estudo
realizado sob a orientação científica dos Doutores Ricardo Manuel Fernandes da Costa (MF-Q,
CEF) e Jorge Manuel Pataca Leal Canhoto (CEF, DCV).
October 2021
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Project “RENATURE - Valorization of the Natural Endogenous Resources of the Centro
Region” (CENTRO-01-0145-FEDER-000007). Funded by the Comissão de Coordenação da
Região Centro (CCDR-C) and subsidized by the European Regional Development Found
(FEDER). Project PTDC/BAA-AGR/32265/2017: “Tamarillo breeding: better plants for better
products”. (project nº 032265 in scope of tender councel nº 02/SAICT/2017). Project Ideas4life
Novel Maritime Plant Food Ingredients (C493053224-00089325); Horizon 2020 Framework
Programme.
Projeto “RENATURE- Valorização dos Recursos Naturais Endógenos da Região do Centro”
(CENTRO-01-0145-FEDER-000007). Fundado pela Comissão de Coordenação da Região
Centro (CCDR-C) e subsidiado pelo Fundo Europeu de Desenvolvimento Regional (FEDER).
Projeto PTDC/BAA-AGR/32265/2017: “Tamarillo breeding: better plants for better products”.
(projeto nº 032265 no âmbito do Aviso de concurso nº 02/SAICT/2017). Projeto Ideas4life
Novel Maritime Plant Food Ingredients (C493053224-00089325); Horizon 2020 Framework
Programme.
Endorsements:
Apoios:
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“Dripping water hollows out stone, not through
force but through persistence.”
Ovid
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VII
ACKNOWLEDGMENTS / AGRADECIMENTOS
Em primeiro lugar saliento que esta dissertação não resulta de um trabalho individual,
mas sim de todo um esforço coletivo. O meu mais sincero obrigado a todos os que me ajudaram
em toda a minha etapa académica e a todos os que riram (e choraram!) comigo.
Um agradecimento especial à minha família direta, esta tese é especialmente vossa. Que
tenham tanto orgulho em mim quanto eu tenho em vocês. Para a minha mãe e pai, obrigado por
todo o apoio, as palavras, os valores e a compreensão. Para a minha irmã e irmão, obrigado por
todos os sermões, todas as brincadeiras e por sempre acreditarem no vosso manito mais novo.
Aos meus orientadores, Dr. Jorge Canhoto e Dr. Ricardo Costa, um sincero obrigado. Por
toda a coordenação e apoio não só académico e prático, mas também moral. Sem a vossa ajuda,
disponibilidade e paciência jamais conseguiria concretizar esta etapa.
Quero agradecer do fundo do meu coração ao João Martins do CEF, por a monumental
ajuda e disponibilidade durante grande parte do trabalho experimental no laboratório, pela
amizade, pela partilha e pelas divertidas conversas que tivemos. Para mim sempre serás o meu
terceiro orientador.
Obrigado à Dr. Sandra Correia por disponibilizar algum do material vegetal utilizado no
estudo, bem como à Dr. Mónica Zuzarte do IBILI por toda a ajuda durante a observação
microscópica do material foliar e sua simpatia.
Deixo também um sincero obrigado a todas as pessoas do CEF e do QMF que de uma
maneira ou de outra me ajudaram durante a parte experimental do trabalho. Obrigado pela vossa
valência e simpatia. A vossa ajuda foi realmente valiosa para a concretização deste trabalho.
Aos meus amigos do coração, que nunca me deixaram cair, que me valorizaram e
apoiaram durante todo este percurso. Abraão Pedra, Alexandre Nogueira, Alexandre Paya,
Carlos Cipriano, Carlos Diogo, Cátia Torres, Diogo Soares, George Lopes, Gonçalo Pereira,
Ivo Costa, Joana Margarido, João Bastos, João Lincho, João Reis, João Silva, João Sousa, Luís
Matos, Luís Rodrigues, Maria Inês Oliveira, Marta Costa, Miguel Dias, Miriam Almeida, Pedro
Pires, Ricardo Carvalho, Ricardo Ferreira, Rita Santos, Sandrine Fernandes, Steve Ferreira,
Thomas Silva, e todos os outros que sei que me esqueci, um obrigado especial meus amores.
Finalmente, quero agradecer a todos os meus colegas de mestrado que me acompanharam
durante os últimos dois anos. Obrigado por toda a ajuda, a companhia e as gargalhadas.
Esta tese não é minha, é de todos nós!
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ABSTRACT
The urge for new environmentally friendly solutions to the pesticides has led to a recent
increasing interest in the research and production of plant-based biopesticides and the
compounds they produce. In the present study, leaves of the species Solanum betaceum Cav.
and Corema album (L.) D. Don were used to create crude ethanol extracts which were
subsequently fractioned using organic solvents. The species leaf fractions were tested in vitro
for their anti-oomycete activity on the growth of Phytophthora cinnamomi Rands. The initial
extract of the leaf material was prepared with ethanol and the fractionation was carried out with
n-hexane, chloroform, ethyl acetate and butanol, in that order, through a biphasic separation
system. P. cinnamomi shows sensitivity to the butanolic fraction of S. betaceum and to all
fractions of C. album, particularly to the initial C. album ethanolic extract. The butanolic
fraction of S. betaceum proves to be rich in polyphenols, polysaccharides and esters. The
various fractions of C. album contain pectins, esters, triterpenoids and aromatic compounds
such as phenols.
Keywords: Anti-oomycete; Corema album; Pesticide; Phytophthora cinnamomi;
Solanum betaceum.
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RESUMO
A necessidade de novas soluções biologicamente sustentáveis ao uso de pesticidas levou
a um recente crescimento no interesse à investigação e produção de biopesticidas à base de
plantas e os compostos por elas produzidos. No presente estudo foram realizados e fracionados
à base de solventes orgânicos frações vegetais de folhas das espécies Solanum betaceum Cav.
e Corema album (L.) D. Don, e a atividade antioomicética dos mesmos foi testada in vitro no
crescimento de Phytophthora cinnamomi Rands. O extrato inicial do material foliar foi
preparado com etanol e o fracionamento do mesmo foi realizado com n-hexano, clorofórmio,
acetato de etilo e butanol, por esta ordem, através de um sistema de separação bifásica. P.
cinnamomi demonstra sensibilidade à fração butanólica de S. betaceum e a todas as frações de
C. album, particularmente ao extrato inicial etanólico. A fração butanólica de S. betaceum
demonstra ser rica em polifenóis, polissacarídeos e ésteres. As várias frações de C. album
possuem pectinas, ésteres, triterpenóides e compostos aromáticos como fenóis.
Palavras-chave: Anti-oomycete; Corema album; Pesticida; Phytophthora cinnamomi;
Solanum betaceum.
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TABLE OF CONTENTS
1. INTRODUCTION ............................................................................................................... 1
1.1. Pesticides ........................................................................................................................ 1
1.2. Phytophthora cinnamomi Rands .................................................................................... 5
1.3. Solanum betaceum Cav. ................................................................................................. 6
1.4. S. betaceum chemical characterization ........................................................................... 8
1.5. Corema album (L.) D. Don .......................................................................................... 14
1.6. C. album chemical characterization ............................................................................. 16
1.7. Trichomes ..................................................................................................................... 18
1.8. Objective....................................................................................................................... 20
2. MATERIALS AND METHODS ...................................................................................... 21
2.1. Plant material ................................................................................................................ 21
2.2. P. cinnamomi cultures and growth medium ................................................................. 22
2.3. Plant extract preparation ............................................................................................... 22
2.4. FTIR –Fourier-Transform Infrared Spectroscopy ........................................................ 27
2.5. Multiwell Activity Assays ............................................................................................ 28
2.6. Extract Fraction Activity Assays .................................................................................. 30
2.7. Trichome observation and scanning electron microscopy ........................................... 32
3. RESULTS ........................................................................................................................... 33
3.1. Multiwell Activity Assays ............................................................................................ 33
3.1.1. Solanum betaceum MAA ...................................................................................... 33
3.1.2. Corema album MAA ............................................................................................. 36
3.1.3. Aliette Flash and DMSO MAA ............................................................................. 38
3.2. Extract Fraction Activity Assays .................................................................................. 39
3.2.1. Control EFAA ....................................................................................................... 39
3.2.2. Solanum betaceum EFAA ..................................................................................... 40
3.2.3. Corema album EFAA ............................................................................................ 42
3.3. FTIR analysis................................................................................................................ 44
3.3.1. Solanum betaceum FTIR analysis ......................................................................... 44
3.3.2. Corema album FTIR analysis ................................................................................ 47
3.4. Trichome identification ................................................................................................ 53
3.4.1. Solanum betaceum trichomes ................................................................................ 53
3.4.2. Corema album trichomes ...................................................................................... 54
4. DISCUSSION ..................................................................................................................... 55
5. CONCLUSION .................................................................................................................. 63
6. REFERENCES .................................................................................................................. 65
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LIST OF ABBREVIATIONS
Abbreviation
Meaning
0-EtOH
Ethanol extract
1-HEX
n-Hexane extract fraction
2-CHL
Chloroform extract fraction
3-ETA
Ethyl acetate extract fraction
4-BUT
n-Butanol extract fraction
5-AQU
Aqueous extract fraction
ATR
Attenuated total reflectance attachment
BE
β-carotene equivalent
C-3-GE
Cyanidin-3-glucoside equivalent
CAM
Portuguese-crowberry (Corema album)
DMSO
Dimethyl sulfoxide
DW
Dry weight
ECHA
European Chemicals Agency
EFAA
Extract activity assay(s)
EFSA
European Food Safety Authority
EW
Edible weight
FA
Fatty acids
FAO
Food and Agriculture Organization of the United Nations
FTIR
Fourier-Transform Infrared Spectroscopy
FW
Fresh weight
GAE
Gallic acid equivalent
GR
Growth reduction
HGA
Hyphal growth area
ISSG
Invasive Species Specialist Group
IU
International unit (1 mg β-carotene = 1667 IU Vitamin A activity)
LBS
Liquid biphasic system
LE
Lutein equivalent
MAA
Multiwell activity assay(s)
MIC
Minimum inhibitory concentration
MLC
Minimum lethal concentration
MUFA
Monounsaturated fatty acids
NAP
National Action Plan
PDA
Potato dextrose agar
PDB
Potato dextrose broth
PES
Polyethersulfone
PTG
Post treatment growth
PUFA
Polyunsaturated fatty acids
QE
Quercetin equivalent
RE
Rutin equivalent
SC-CO2
Supercritical CO2 extraction
SE
Soxhlet extraction
SEM
Scanning electron microscopy
SFA
Saturated fatty acids
TAM
Tamarillo (Solanum betaceum)
WHO
World Health Organization
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1. INTRODUCTION
1.1. Pesticides
Pesticides - Definition and their applications
A pesticide is a chemical substance, or a biological agent intentionally released into the
environment, capable of deterring, preventing or controlling populations of harmful pests such
as animals, weeds, fungi, bacteria or viruses. These pests are defined as organisms that can be
hazardous to our food and health (Mahmood et al., 2016) and are liable for yearly expenses of
billions of dollars through the production of costly synthetic chemicals (Chattopadhyay et al.,
2017). The term “pesticide” is a general term that refers to a wide range of compounds such as
insecticides, fungicides, herbicides, rodenticides, molluscicides, nematicides, among others.
Insecticides are generally the most acutely toxic class of pesticides (Aktar et al., 2009).
The world population is largely dependent of agrochemicals use, being the only reliable
way of protecting large scale food production (Andréa et al., 2000) necessary for the subsistence
of an ever-growing world population. That being said, the use of pesticides seems to be a
necessary evil to mankind food necessities as a whole (Carvalho, 2006; Peshin & Zhang, 2014).
This assessment is supported by the increasing production and use of these chemicals around
the world (Fig. 1).
Figure 1. Average use of pesticides per area of cropland worldwide from 1990-2019.
(source: fao.org/faostat. Accessed: 12/08/2021)
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Pesticides polution and risks
Pesticides overuse and pollution is increasing around the globe, especially in countries
outside Europe, such as India, China and Brazil (Zhang & Liu, 2017). The frequent use of
pesticides negatively affects soils and water bodies quality, and contaminate surface waters by
drift, run-off, drainage and leeching (Houtman, 2010). Studies conducted in Italy reveal
dangerously high concentrations of pesticides in surface and groundwaters in most part
associated with agricultural activities taken place at local watersheds (Meffe & de Bustamante,
2014). Other monitoring studies conducted in water bodies around the world have shown
similar concerning results (Gao et al., 2009; Fadaei et al., 2012; De Gerónimo et al., 2014).
Being designed to be biologically active (Schuster & Schröder, 1990a), pesticides can affect
other species causing unintended side effects and accumulate on crops that enter our food chain
and are directly ingested by humans along foodstuffs and water (Taylor et al., 2002;
FAO/WHO, 2006). Besides ingestion, two other main routes of human body exposure to
pesticides are inhalation and dermal exposure (Tomer et al., 2015). The exposure to these
agrochemicals is responsible for numerous cases of related illnesses and injuries, especially
among agricultural industry workers, who are directly exposed to these pesticides. Data reports
by Calvert et al. (2016) in the USA reveals that the rates of pesticide-related ailments are
staggeringly greater among agricultural workers that in nonagricultural workers.
The general population is also directly exposed to agrochemicals, mainly in public places
such as office buildings, restaurants, schools, parks, and along roads and walkways (Bolognesi
& Merlo, 2019). Pesticide exposure may result in biochemical alterations in the body long
before more glaring and adverse symptoms are manifested (Tomer et al., 2015). The chronic
exposure to pesticides significantly increase the risk of developing various pathophysiological,
respiratory and neurological conditions (Agrawal & Sharma, 2010). For example, the natural
occurring rotenone (derived from plant roots), widely used as an active agent in insecticides
commonly applied in gardens, lakes and reservoirs, is proven to induce behavioral and
neuropathological features of Parkinson’s disease (Panov et al., 2005; Agrawal & Sharma,
2010). Given the hazardous nature of synthetic pesticides, rigid regulations are imposed to their
use and many known substances are regulated and prohibited, especially at an European Union
(EU) level and its Member States.
3
Pesticide use and legislation in the EU
There are strict legislations concerning the use of pesticides in EU, which aim to regulate
all types of plant protection products and biocidal usage across all EU Member States. Pesticide
regulations are undertaken at an EU level by Parliament directives alongside Member States
national laws, or National Action Plans (NAPs). Although NAPs have the freedom to
implement their particular laws towards the use of pesticides within the given country borders,
these regulations must comply and work in tandem with the collective EU. NAPs should
advocate a proper training amongst producers, distributors, advisors and professional users of
pesticides, the implementation of certification systems explicitly disclaiming potential health
and environmental risks, and the public sensibilization and education towards the use of
agrochemicals (Directive 2009/128/EC of the European Parliament and of the Council of 21
October 2009). Pesticide placement in the market is also thoroughly revised. These regulations
are mediated the Regulation on Plant Protection Products (Regulation No 1107/2009 of the
European Parliament and of the Council, 2009) and the Regulation on Biocidal Products
(Regulation No 524/2013 of the European Parliament and of the Council, 2012). Those
regulations were implemented with the objective of protecting human, animal and environment
health, as well as to standardize the rules to which the plant protection products and biocides
must adhere to be approved for marketing. Both types of agrochemicals are subject to a dual
approval process, where the products active substances are revised at an EU level and the
products themselves are at a Member State level. The approval of active substances by the EU
is mediated by the European Food Safety Authority (EFSA) and the European Chemicals
Agency (ECHA) by a set number of “exemption criteria” (Table 1). As of June 2021, EU
regulation entities assesses the state of 1518 active substances, among them 456 are approved,
921 are not approved, 61 are under review and 17 are yet to be assessed at an EU level.
Plant protection substances
Biocidal substances
Effects on
human health
Cannot be classified as carcinogenic, mutagenic or toxic to reproduction
Cannot be considered an endocrine disruptor
Effects on the
environment
Cannot be considered a persistent
organic pollutant (POP)
Cannot be considered as a persistent, bio-accumulative and toxic (PBT)
substance, or a very persistent, very bio-accumulative substance (vPvB)
Table 1. Principal exemption criteria for substance approval in the EU.
(Adapted from European Parliamentary Research Service 2017)
NOTE: EU is referring to European Union, which excludes all non-Member States as of June 2021.
4
Biopesticides and alternatives
Given the potentially hazardous consequences of pesticide overuse and the strict
regulations imposed in the pesticides active substances and products, it is of utmost importance
to search for environmentally friendly options in our immediate future. The use of organic-
based compounds seems to be a potential solution to develop effective new products to tackle
pest management. This biopesticides, as they are named, have several crucial differences that
distinguishes them from chemical pesticides. Their active compounds come from animals,
plants and bacteria (EPA, n.d.) and usually target specific pests and leave the remaining
organisms unscathed (Dar et al., 2021). They are also biodegradable with little to no residual
effects and are less prone to be rendered useless by pest resistances (Gupta & Dikshit, 2010).
Currently there are many plant-based products in use in organic agriculture, with effective
results in the control of many economical important pests, such as biopesticide products based
of neem (Azadirachta indica) (Khater, 2012) and butterfly-pea (Clitoria ternatea) (Damalas &
Koutroubas, 2018).
Pesticides and Phytophthora cinnamomi
Currently, only a single class of pesticides has proven to be effective in the control of the
pathogen P. cinnamomi and readily available in the global market, the phosphites. This class of
compounds are anionic forms of phosphonic acid (H3PO3) and their application in the infected
plant decrease (but not prevent) the production of the oomycete sporangia and zoospores, and
trigger a defense response in the targeted plant which inhibit the oomycete growth (Guest &
Grant, 1991; Wilkinson et al., 2001). Despite these positive effects, zoospores produced by P.
cinnamomi previously sprayed with phosphites are still viable and capable of infecting new
plants (Wilkinson et al., 2001). Phosphites are also phytotoxic, inducing foliar necrosis in many
studied species. Their intake and retention is highly variable from species to species and is
directly correlated with the appearance and severity of the symptoms shown in treated plants,
making these compounds application potentially harmful to some species (Pilbeam et al., 2000;
Barrett et al., 2004). The practical application of these phosphites in large areas is conducted
via aerial spraying, covering the entirety of the treated areas that usually comprises communities
with rare and threatened plant species (Hardy et al., 2001). These detrimental consequences that
come from the use of pesticides is unfortunately very common and does not only extend to the
destruction of the surrounding flora, but also the contamination of the environment soils and
waters.
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1.2. Phytophthora cinnamomi Rands
P. cinnamomi is a diploid pathogenic oomycete (Kroon et al., 2004) capable of infecting
a large number of plant species (likely close to 5000 species), among them important crops such
as the avocado, chestnut, peach and pineapple (Hardham, 2005). The pathogen was first
described by Rands in the west coast of Sumatra, causing bark canker in infected cinnamon
trees (Cinnamomum burmanni), hence its name (Rands, 1922). P. cinnamomi exhibits mycelial
growth and produces motile flagellated asexual zoospores (Hardham, 2005; Beakes et al.,
2012). The pathogen has both asexual and sexual phases during its life cycle, being the former
favored over the latter (Hardham, 2005) (Fig. 2).
This oomycete has a cosmopolitan global distribution (Fig. 3), reproducing rapidly in
moist soils and in temperatures of 16-30 ºC, ideally around 22-28 ºC, at 4.0-7.0 pH (Chee &
Newhook, 1965) and can survive up to six years at favorable conditions (Zentmyer & Mircetich,
1966). Thriving in waterlogged soils and after events of heavy rain, moisture is crucial to the
development of the sporangia and the production and movement of the motile zoospores
(Hardham, 2005). The oomycete is a major threat to natural ecosystems and biodiversity
worldwide, causing heavy economic losses in agriculture and forestry (Hardham & Blackman,
2018). The pathogen causes the rotting of fibrous roots and stem cankers, as well as the dieback
of young shoots. The oomycete usually invades its host via the finer roots, where the zoospores
form cysts that germinate and grow a tube that penetrates the outer cell layer (Cahill et al.,
2008). The infected roots have a significantly reduced water intake, which results in foliage
chlorosis and the rapid wilting of the plant (Hardham & Blackman, 2018).
Figure 2. Phytophthora cinnamomi. (A) Asexual spore, sporangia; (B) Sexual spore, oogonia
and antheridia.
(source: bugwood.org. photos: A - Elizabeth Bush; B - Mary Ann Hansen)
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Having a large number of possible hosts, P. cinnamomi is responsible for many types of
plant diseases worldwide, such as the Jarrah Dieback (Eucalyptus marginata) in south-western
Australia (Shearer & Tippett, 1989; Hardham, 2005), root rot in Quercus species in the Iberian
Peninsula (Brasier et al., 1993; Brasier, 1996), stem canker in peach tress (Prunus persica) in
several USA states (Haygood et al., 1986; Mircetich & Keil, 1970) and dieback or “ink disease”
in chestnut trees (Castanea sativa) (Crandall et al., 1945; Vettraino et al., 2005) across Europe,
among others. The oomycete also indirectly affects the fauna of the habitats it infects, disrupting
food chains and reducing birds nesting sites, as the infection results in the substantial loss of
plant canopy area (Cahill et al., 2008; Garkaklis et al., 2004). Given its high contagion rate and
global dispersion, the pathogen was placed in the top 100 most dangerous species in the world
by the Invasive Species Specialist Group (ISSG) (van der Weijden et al., 2017).
No thoroughly effective solutions currently exist to eradicate P. cinnamomi once the
infection is established. Human activity management is the main approach to P. cinnamomi
control, as humans are the fastest and widest vector of this oomycete propagation, which by
itself spreads rather slowly (O’Gara et al., 2005). This management consists in the adoption of
preventive practices and operations in risk-prone areas and field personnel sensibilization and
training (Hardy et al., 2001).
1.3. Solanum betaceum Cav.
Taxonomy, ecology and distribution
Solanum betaceum (Cav.), tamarillo or “tree-tomato” (Wiersema & León, 2016) belongs
to the Solanum genus and Solanaceae family. This genus accounts for half this family diversity
Figure 3. Global distribution of Phytophthora cinnamomi.
(source: cabi.org. Accessed: 17/06/2021)
7
and is considered the most economically important genus within Solanaceae (Frodin, 2004;
Bohs, 2007). The species is a subtropical plant native to the Andes of Peru, Chile, Ecuador,
Colombia and Bolivia (Fig. 4A), where wild populations of tamarillo occur along with the other
closely related species of the same genus (Bohs, 1989b; Prohens & Nuez, 2001). The tamarillo
tree is commonly found growing in mountainous regions in medium to high altitudes, especially
in its native regions of South America. (Morton, 1982; Duke & DuCellier, 1993). Ideally,
tamarillo naturally grows in regions with an average annual temperature of 15 to 25 ºC, and
rainfall between 1000-2000 mm (minimum 800 mm). Tamarillo flowers and fruits throughout
the entire year, suggesting no photoperiodic response (Duarte & Paull, 2015). The species was
probably spread by the Spanish throughout Latin America, and later was dispersed to the tropics
and subtropics of Africa, South Asia, China, Australia, New Zealand, USA and Europe (Fig.
4B). Studies report that the tamarillo shows effective establishment conditions in the
Mediterranean region (Prohens et al., 1997). To this day the species is cultivated in all these
regions around the world, such as Jamaica, Puerto Rico, Costa Rica, Haiti, Guatemala, Sri
Lanka, New Guinea, China, South Africa and many others (Bohs, 1989b; Duke & duCellier,
1993).
Botanical characterization
S. betaceum is a tree which normally grows 2-4 m tall. Its branches are narrow and brittle,
branching out of the main stem to form a wide dichasial crown (Bohs, 1989b; Bohs, 1994;
Lewis & Considine, 1999b) (Fig. 5A). The tree starts to produce fruits after 1-2 years (Prohens
Figure 4. Distribution of S. betaceum in the native region of South America. (A) Native distribution
in South America; (B) Global dispersal. (Adapted from Bohs, 1989b)
8
& Nuez, 2001) and can produce fruit for 8-12 years (Bakshi et al., 2016). The tamarillo tree has
large simple leaves with long petioles, are softly pubescent on both sides and have a
characteristic musky smell. (Bohs, 1989b; Bohs, 1994; Prohens & Nuez, 2001; Duarte & Paull,
2015). S. betaceum flowers grow in buds of 10-50 flowers (Lewis & Considine, 1999b). They
grow 1.5-2 cm wide, have a stellate shape with a pinkish-white pentamerous corolla, green-
purple calyx and five thick stamens. Tamarillo flower petals are fleshy, glabrous and slightly
curved at the tip (Bohs, 1994; Lewis & Considine, 1999b; Duarte & Paull, 2015) (Fig. 5B).
The fruit has a ellipsoid to ovoid shape and grow singly or in clusters of 3-12 fruit sets
(Bohs, 1994; Duarte and Paull, 2015). They are glabrous with a yellow to orange, red or purple
color, often presenting long longitudinal stripes depending on the fruit variety (Fig. 5C). The
mesocarp is meaty and firm, with a bland to bitter flavor (Bohs, 1989b; Bohs, 1994; Duarte &
Paull, 2015). The fruit grows rapidly, reaching full size in about 16 weeks and maturity in
around 27 weeks after anthesis (Heatherbell et al., 1982). The fruit is acidic with a pH=4.0 ±
0.17 (Romero-Rodriguez et al., 1994) and is highly nutritious (Morton, 1982).
1.4. Solanum betaceum chemical characterization
Phenolic compounds
Having many varieties and cultivars, tamarillo bioactive compounds concentration differ
between varieties and cultivars (Acosta-Quezada et al., 2015). Nevertheless, all tamarillo fruit
varieties present high concentrations of similar phenolic compounds, conferring the fruit
Figure 5. Solanum betaceum. (A) Young tamarillo tree supported by a stake; (B) Tamarillo flower
bud with flowers in different stages of maturation; (C) Tamarillo fruit (red-type).
(photos: A - Bruno Costa; B,C – Gonçalo Pereira)
9
substantial antioxidant properties (Acosta-Quezada et al., 2015; Espin et al., 2016). Ghosal et
al. (2013) reports that the fruit accumulates phenols as it matures, reaching an high
concentration of these compounds in later maturity stages. This increasingly higher reducing
power throughout fruit ripening was also observed by Vasco et al. (2009). Studies in cultivars
from Spain (Espin et al., 2016) and Ecuador (Vasco et al., 2009) report hydroxycinnamoyl
derivatives (yellow-giant: 60.25-110.23 mg/100 g DW; purple-giant: 132.57-421.55 mg/100 g
DW) and hydroxycinnamic acid derivatives (golden-yellow: 39 mg/100 g FW; purple-red: 61
mg/100 g FW) as the most abundant phenolic compounds present in the fruits, respectively.
Mertz et al. (2009) also reports an abundant presence of hydroxycinnamic acids in S. betaceum
fruit extracts, such as caffeoylquinic acid (red: 54.8 mg/100 g DW; yellow: 32.8 mg/100 g DW).
DW stands for “dry weight”. Tamarillo phenolic content is found in the Table 2.
Anthocyanins and flavonoids
Tamarillo fruits presents a variable concentration of anthocyanins according to cultivar
location and fruit color (Hurtado et al., 2009; Espin et al., 2016). These compounds accumulate
as the fruit ripens (Heatherbell et al., 1982) and are more concentrated in the fruit pulp (4.15
mg C-3-GE/100 g DW) than in the peel (1.36 mg C-3-GE/100 g DW) (Hassan & Bakar, 2013).
The yellow fruit varieties tend to have negligeable amounts of anthocyanins, as shown in Vasco
et al. (2009), Mertz et al. (2009) and Espin et al. (2016). The major anthocyanins found in
tamarillo fruit are delphinidin-3-rutinoside, cyanidin-3-rutinoside and pelargonidin-3-
rutinoside (Wrolstad & Heatherbell, 1974; Mertz et al., 2009; Hassan & Bakar, 2013).
Delphinidin-3-rutinoside is the main anthocyanin in the fruit pulp and cyanidin-3-rutinoside is
the main anthocyanin in the fruit peel, in most of the worldwide studied varieties. Pelargonidin-
3-rutinoside is the main anthocyanin present in studied Ecuadorian cultivars (Wrolstad &
Heatherbell, 1974). The total flavonoid content was higher in the fruit peel (3.36 mg RE/g) than
in the fruit pulp (2.41 mg RE/g) (Hassan & Bakar, 2013). Vasco et al. (2009) found the flavonols
quercetin (golden-yellow: 6 mg/100 g FW; purple-red: 4 mg/100 g FW) and myricetin (golden-
yellow: 1.2 mg/100 g FW; purple-red: 1.4 mg/100 g FW) in tamarillo fruits. FW and RE stands
for “fresh weight” and “rutin equivalent”, respectively. Tamarillo anthocyanin and flavonoid
content is found in the Table 2.
10
Fatty acids
S. betaceum fruits are extremely poor in lipids (Wang & Zhu, 2020). Previous studies by
Castro-Vargas et al. (2013) report low yields of nonpolar lipidic compounds in the pericarp of
tamarillo fruits, corroborating this assessment to a certain degree. Ramakrishnan et al. (2013)
and Achicanoy et al. (2018) conducted studies assessing the fatty acid composition of tamarillo
seeds using supercritical CO2 extraction (SC-CO2) and Soxhlet extraction (SE). Both studies
revealed that tamarillo seeds seem to be much richer in fatty acids than the rest of the fruit, as
proven by lipid extraction yields: 17.4%, optimal SC-CO2 yield; 21.13 and -24.05%, SE yield.
The major type of fatty acids (FA) present in the fruit seeds are polyunsaturated fatty acids
(PUFAs) (72.20% and 72.05%), followed by monounsaturated fatty acids (MUFAs) (15.47%
and 16.18%), and saturated fatty acids (SFAs) (12.32% and 11.80%). The most abundant fatty
acids in tamarillo seeds are linoleic (SC-CO2: 66.67%; SE: 70.47% and 71.30%), oleic (SC-
CO2: 17.94%; SE: 14.93% and 17.3%) and palmitic (SC-CO2: 10.41%; SE: 9.41% and 8.0%)
(Ramakrishnan et al., 2013; Achicanoy et al., 2018). Tamarillo fatty acids content is found in
the Table 3.
Reference
[A]
[B]
[C]
[D]
Region
Ecuador
Penampang, Malaysia
New Zealand
Loja, Ecuador
Obs.
Golden-yellow
Purple-red
Striped-red
N.D.
Orange +
orange-pointed
Red +
red-pointed
Purple
Pulp
Peel
Pulp
Peel
Total phenolic compounds
125 ± 6.21
187 ± 3.71
261 ± 12.03
489 ± 4.03
-
-
2.43 – 6.186
2.94 – 4.396
2.58 – 6.126
Anthocyanins
undetected
38 ± 0.22
2.15 ± 0.144
1.36 ± 0.14
0.987
0.327
-
-
-
Flavonoids
-
-
241 ± 2.05
336 ± 1.05
-
-
-
-
-
[A] Vasco et al., 2009; [B] Hassan & Bakar, 2013; [C] Wrolstad & Heatherbell, 1974; [D] Acosta-Quezada et al., 2015.
1 Gallic acid equivalent (mg GAE/100 g FW)
2 Cyanidin-3-glucoside equivalent (mg C-3-GE/100 g FW)
3 Gallic acid equivalent (mg GAE/100 g DW); Results converted from mg GAE/g to mg GAE/100 g
4 Cyanidin-3-glucoside equivalent (mg C-3-GE/100g DW)
5 Rutin equivalent (mg RE/100 g DW); Results converted from mg RE/g to mg RE/100 g
6 g/100 g DW
7 μmol/g FW
Reference
[A]
[B]
Region
Cameron Highlands, Malaysia
Colombia
Obs.
Seed oil (%)
Seed oil (%)
Polyunsaturated fatty acids
72.20
72.05
Monounsaturated fatty acids
15.47
16.18
Saturated fatty acids
12.32
11.80
[A] Ramakrishnan et al., 2013; [B] Achicanoy et al., 2018.
Table 2. Phenolic compounds content in S. betaceum fruits.
Table 3. Fatty acid percentage content in S. betaceum seed oil.
11
Carotenoids
Tamarillo has a high content in terpenoids (Wang & Zhu, 2020), mainly in carotenoids.
Studied cultivars from Ecuador (Mertz et al., 2009), Colombia (Giuffrida et al., 2018), Brazil
(Rodriguez-Amaya et al., 1983) and China (Yang & Zhao, 2013) present a rich diversity of
carotenoids, such as β-carotene, cryptoxanthin, lutein and zeaxanthin. The commonest
carotenoids found in the tamarillo fruit are precursors of provitamin A (Bauernfeind, 1972),
making the fruit a valuable source of vitamin A (Rodriguez-Amaya, 1999). Rodriguez-Amaya
et al. (1983) reports a vitamin A value of 2475 IU/100 g EW, and Mertz et al. (2009) 2000
RE/kg FW. EW stands for “edible weight”. Hassan & Bakar (2013) determined that the total
carotenoid content in the pulp is 25.13 mg BE/100g DW, and 19.13 mg BE/100 g DW in the
peel. Rodriguez-Amaya et al. (1983) report similar carotenoid concentration results, with 24.3
μg/g FW in the pulp and 22.0 μg/g FW in the peel. “BE” stands for “β-carotene equivalent”.
Studies in extracts of tamarillo fruit indicate that the major free carotenoids in tamarillo fruits
are β-carotene and β-cryptoxanthin. (Rodriguez-Amaya et al., 1983; Mertz et al., 2009;
Giuffrida et al., 2018) Carotenoid esters account for 78% of the total carotenoid composition
(Mertz et al., 2009). Tamarillo carotenoids content is found in the Table 4.
Alkaloids
The presence of alkaloids and other N-containing compounds in S. betaceum is well
documented. Nevertheless, the specific plants organs where these compounds exist and their
quantification is not clearly reported (Wang & Zhu, 2020). Eich (2008) compiled and described
various alkaloids found in S. betaceum, alongside the studies in which these compounds were
found. Pyrrolidines like solamine, solacaproine, tropinone, cuscohygrine and tomatidenol have
Reference
[A]
[B]
[C]
Region
Ecuador
Brazil
Penampang, Malaysia
Obs.
Yellow
(μg LE/g FW)
Red
(μg LE/g FW)
Fruit pulp
(μg/g FW)
Fruit peel
(μg/g FW)
Fruit pulp
(mg BE/100 g DW)
Fruit peel
(mg BE/100 g DW)
β-Carotene
4.6 ± 0.31
5.1 ± 0.31
7.9 ± 3.6
8.8 ± 3.5
-
-
Cryptoxanthin
1.1 ± 0.11
1.5 ± 0.081
13.9 ± 4.2
10.0 ± 2.8
-
-
Zeaxanthin
0.1 ± 0.021
0.3 ± 0.061
0.6 ± 0.6
1.1 ± 0.6
-
-
Lutein
0.98 ± 0.052
1.25 ± 0.052
1.7 ± 1.1
1.5 ± 0.3
-
-
Total carotenoids
-
-
24.3
22.0
25.13 ± 0.35
19.13 ± 1.93
[A] Metz et al., 2009; [B] Rodriguez-Amaya et al., 1982; [C] Hassan & Bakar, 2013.
1 Before saponification; 2 After saponification (Lutein concentration is exceptionally noted in β-carotene equivalents (μg BE/g FW).
Table 4. Total carotenoid content and main carotenoids in S. betaceum fruits.
12
been found in unspecified parts of the tamarillo plant (Wang & Zhu, 2020). Evans et al. (1972)
also reports the presence of the aforementioned compounds (with the exception of tomatidenol)
in the tamarillo roots, alongside other compounds like amines. Schröter & Neumann (1964)
also describe the presence of tropane and steroid alkaloids in S. betaceum roots. Trace amounts
of tropane-derived calystegins were found in tamarillo fruits by Asano et al. (1997b), but more
recent tests seem to indicate the absence or an undetectable concentration of alkaloids in S.
betaceum fruits (Vasco et al., 2009).
Sugars and other acids
Tamarillo fruits have a moderate amount of sugar compounds and other carbohydrates
such as organics acids and pectins. In comparison with other tropical fruits, S. betaceum holds
a relatively low sugar content (Wills et al., 1986; Vasco et al., 2009), which make the fruit an
healthier alternative to its counterparts. The sugar concentration increases as the fruit matures,
and it slightly varies between different tamarillo varieties (Heatherbell et al., 1982; Acosta-
Quezada et al., 2015). The three main sugars found in the tamarillo fruit are glucose, sucrose
and fructose (Heatherbell et al., 1982). S. betaceum fruits present considerable amounts of citric
acid and ascorbic acid (vitamin C), with comparatively low concentrations of malic acid and
quinic acid (Romero-Rodriguez et al., 1994; Vasco et al., 2009; Acosta-Quezada et al., 2015).
Both citric acid and malic acid are produced during fruit maturation, and their concentration
decreases substantially after fruit ripening (Heatherbell et al., 1982). Various monosaccharides
were isolated from tamarillo pulp and seed mucilage, the main ones being arabinose, xylose,
galactose and uronic acids. The detection of these compounds indicate the presence of long
chain pectins in the tamarillo fruit and mucilage (do Nascimento et al., 2013; do Nascimento et
al., 2016). Cacioppo (1984) and Romero-Rodriguez et al. (1994) report a high content of
ascorbic acid in yellow (30.0-35.0 mg/100g FW and 19.7 mg/100g FW, respectively) and red
(35.0-45.0 mg/100g FW and 21.9 mg/100g FW, respectively) tamarillo varieties, making its
fruits a great source of vitamin C. Tamarillo sugar and acid content is found in the Table 5.
13
Minerals
The mineral content in tamarillo fruits is highly variable between different cultivars and
regions, but all studied cultivars are particularly rich in potassium and phosphorus, minerals
which are usually present at low concentrations in fruits (Dawes & Callaghan, 1970). Romero-
Rodriguez et al. (1994) reports concentrations of potassium as high as 404 ± 47.1 mg/100 g FW
in yellow tamarillo variant in Galicia cultivars and Vasco et al. (2009) reports concentrations
reaching 398 ± 11.3 mg/100 g FW in Ecuador cultivars. The species fruits also hold a relatively
high concentration of calcium and magnesium (Vasco et al., 2009; Acosta-Quezada et al.,
2015). Tamarillo mineral content is found in the Table 6.
Reference
[A]
[B]
[C]
[D]
Region
Galicia, Spain
Ecuador
Loja, Ecuador
New Zealand
Obs.
Yellow
(%)
Red
(%)
Golden-yellow
(%)
Purple-red
(%)
Orange +
orange-pointed
(g/100 g DW)
Red +
red-conical
(g/100 g DW)
Purple
(g/100 g DW)
ASR
(g/100 g FW)
SR
(g/100 g FW)
GM
(g/100 g FW)
Glucose
0.5 ± 0.10
1.0 ± 0.23
1.7 ± 0.02
1.4 ± 0.10
6.3 – 12.4
8.0 – 10.2
7.4 – 9.6
0.774
0.707
0.923
Fructose
0.7 ± 0.14
1.2 ± 0.14
1.6 ± 0.10
1.4 ± 0.10
6.3 – 13.2
8.4 – 9.9
7.4 – 10.1
1.079
0.910
1.139
Sucrose
1.6 ± 0.26
2.5 ± 0.27
1.9 ± 0.10
1.7 ± 0.10
15.5 - 27.1
20.8 – 24.1
18.3 – 25.8
1.813
1.100
1.891
Citric acid
1.8 ± 0.11
1.7 ± 0.03
2.5 ± 0.10
2.7 ± 0.04
5.42 – 7.54
4.01 – 6.39
5.50 – 6.91
1.27
1.35
1.71
Malic acid
0.07 ± 0.01
0.05 ± 0.01
0.07 ± 0.03
0.53 ± 0.02
0.34 – 0.78
0.46 – 0.99
0.53 – 0.88
0.14
0.13
0.15
Quinic acid
0.8 ± 0.06
0.4 ± 0.03
-
-
-
-
-
-
-
-
[A] Romero-Rodriguez et al., 1994; [B] Vasco et al., 2019; [C] Acosta-Quezada et al., 2015; [D] Boyes & Strübi, 2010.
ASR – Andys Sweet Red variety; SR – Secombes Red variety; GM – Goldmine variety.
Reference
[A]
[B]
[C]
[D]
Region
Galicia, Spain
Ecuador
New Zealand
Loja, Ecuador
Obs.
Yellow
(mg/100 g FW)
Red
(mg/100 g FW)
Golden-yellow
(mg/100 g FW)
Purple-red
(mg/100 g FW)
Yellow
(mg/100 g FW)
Red
(mg/100 g FW)
Orange +
orange-pointed
(mg/100 g DW)
Red +
red-conical
(mg/100 g DW)
Purple
(mg/100 g DW)
Sodium (Na)
4.9 ± 0.66
8.9 ± 2.9
0.06 ± 0.001
0.20 ± 0.001
-
-
-
-
-
Potassium (K)
404 ± 47.1
347 ± 14.7
398 ± 11.3
379 ± 3.4
292.0
321.0
-
-
-
Calcium (Ca)
10.6 ± 0.05
9.3 ± 1.09
25 ± 0.8
22 ± 0.2
11.0
11.0
34 – 80
42 – 70
16 – 78
Magnesium (Mg)
22.3 ± 2.09
19.7 ± 1.8
16 ± 0.5
14 ± 0.4
20.0
21.0
58 – 230
48 – 221
54 – 214
Iron (Fe)
0.4 ± 0.03
0.4 ± 0.05
0.22 ± 0.02
0.46 ± 0.01
0.44
0.57
0.68 – 2.32
1.12 – 1.64
1.00 – 2.43
Copper (Cu)
0.2 ± 0.06
0.2 ± 0.05
0.08 ± 0.01
0.12 ± 0.01
0.06
0.05
0.07 – 3.25
0.06 – 0.82
0.07 – 0.70
Zinc (Zn)
0.2
0.2 ± 0.03
0.20 ± 0.02
0.17 ± 0.01
0.17
0.15
0.21 – 1.44
0.46 – 1.83
0.35 – 1.13
Manganese (Mn)
0.1
0.1
-
-
0.185
0.114
-
-
-
Phosphorus (P)
-
-
-
-
40.0
39.0
-
-
-
[A] Romero-Rodriguez et al., 1994; [B] Vasco et al., 2009; [C] Diep et al., 2020; [D] Acosta-Quezada et al., 2015.
Table 5. Sugar and organic acids compounds in S. betaceum fruits.
Table 6. Mineral content in S. betaceum fruits.
14
1.5. Corema album (L.) D. Don
Taxonomy, ecology and distribution
Corema album (L.) D. Don, Portuguese-crowberry or “camarinha” is a member of the
Ericaceae family. Corema is a small genus with only two species, C. album and C. conradii,
both being small coastal dioecious shrubs with drupaceous fruits. C. album has two subspecies,
C. album subsp. album and C. album subsp. azoricum P. Silva, the latter being endemic to the
Azores islands of Faial, Graciosa, Pico, São Jorge and São Miguel (DRAAC, 2010), being a
protected species in the region (Decreto Legislativo Regional 15/2012/A of April 2 of Regime
Jurídico Da Conservação Da Natureza e Da Proteção Da Biodiversidade, 2012).
C. album occurs in the Atlantic coast of the Iberian
peninsula (León-González et al., 2013). The crowberry
habits sand dunes and rocky-coast sites (Guitián et al.,
1997). Its distribution spans the entire west Iberian
peninsula coastal region, from the north of Galicia to the
south of Gibraltar (Álvarez-Cansino et al., 2010;
Zunzunegui et al., 2006), except for the coastal region of
Douro in Portugal, where the species does not seem to be
established (Blanca et al., 2000) (Fig. 6). Crowberry is
commonly found forming associations alongside species
like Pinus pinea, Juniperus phoenicea, Pistacia
lentiscus, Calluna vulgaris, Cistus crispus, among
several other species (Ferreira, 2018; López-Dóriga,
2018). Crowberry is found from seaside up to 50 meters
above sea level (López-Dóriga, 2018), growing in
regions with an annual temperature of roughly 15 ºC
(13.4-16.8 ºC) and high variations in annual rainfall,
ranging from 540 mm to 1355 mm, according to reports
from studied population sites in Álvarez-Cansino et al. (2013). C. album can withstand harsh
summer drought events, which are progressively common in the Mediterranean basin
(Christensen & Christensen, 2007). The species is especially vulnerable to habitat changes, as
it inhabits coastal regions prone to touristic recreation, infrastructure expansion and sand
gathering for construction materials (Blanca et al., 2000; Clavijo et al., 2002; Gil-López, 2011).
Figure 6. Distribution of C. album
in the Iberian Peninsula coastline,
with several studied populations
marked. (Adapted from Álvarez-
Cansino et al., 2013)
NOTE: Although the name “crowberry” is more commonly used to identify the Empetrum
nigrum species (black-crowberry), throughout the entire thesis Corema album (Portuguese-
crowberry) is commonly referred as “crowberry” for readability sake.
15
Currently in Spain there are only two populations of C. album that have male and female plants
in all age ranges and therefore relatively safe from extinction, while all the other isolated
populations of crowberry consist mainly of old individuals and are doomed to disappear, unless
protective conservation measures are urgently implemented (Aguilella & Laguna, 2009).
Botanical characterization
C. album is a perennial evergreen shrub that grows between 30–75 cm tall (Guitián et al.,
1997; Oliveira & Dale, 2012), is densely branched from the base and can reach up to 3 m in
diameter (Clavijo et al., 2002) (Fig. 7A). Although the crowberry is mostly a dioecious species,
hermaphrodite crowberries may occur in low proportions, only occurring in bigger ratios in
specific regions and populations (Zunzunegui et al., 2006). The species deep root system can
be complemented by adventitious roots sprouting from branches that may become buried in the
sediment, such as sand blown by the wind (Álvarez-Cansino et al., 2010). C. album has small
linear ericoid dark-green leaves with a prominent groove along the abaxial axis. This groove is
a result of the leaf blade folding into itself (Tutin et al., 1972; Villar, 1993; Oliveira & Dale,
2012). Crowberry leaves have a short petiole and grow laid against the stem alternated in whorls
of 3-4 leaves (Villar, 1993; Oliveira & Dale, 2012). The leaves are covered in sessile glands
when young, becoming glabrous when mature (Tutin et al., 1972).
C. album flowers are tightly packed in terminal racemose inflorescences of variable
morphology, depending on the plant sex (Guitián et al., 1997; Oliveira & Dale, 2012). Female
plants have 1-2 flowers per inflorescence and male plants have 4-14 flowers per inflorescence
with ovate to acuminate bracts (Villar, 1993; Zunzunegui et al., 2006). Both the male and
female flowers are actinomorphic with 3 pinkish-red obovate petals and 3 green pubescent
sepals (Fig. 7B, 7D). Male flowers have 3 exert stamens with conspicuous red-purple anthers
(Villar, 1993; Oliveira & Dale, 2012). Female flowers are smaller than their male counterparts,
and in some cases may completely lack their petals (Tutin et al., 1972). Crowberries usually
flowers from early March to mid-April, but can flower as early as February (Blanca et al., 2000).
The fruit is a small pinkish-white spherical berry-like drupe (Tutin et al., 1972; Guitián et al.,
1997; Oliveira & Dale, 2012) (Fig. 7C), highly acidic with a pH of around 3, a moisture content
of 81.7% (Brito et al., 2021) and a characteristic lemony flavor. The fruit usually contains three
seeds enclosed by a thick woody endocarp (pyrenes), but the number can change between 2-9
(Guitián et al., 1997; Calviño-Cancela, 2002; Zunzunegui et al., 2006). Fruiting occurs during
early summer through late autumn, peaking during August and early September (Oliveira &
Dale, 2012).
16
1.6. Corema album chemical characterization
Phenolic compounds
C. album is rich in phenolic compounds, especially in the plant leaves and fruits,
conferring the plant a high antioxidative power (Brito et al., 2021). Crowberry fruits have a
high concentration of phenolic acids (2268.1 mg/kg DW), flavonoids (1437 mg GAE/100 g
pulp), tannins (871 mg GAE/100 g pulp), and in lower concentrations, ortho-diphenols (21.8
mg GAE/100 g pulp) and anthocyanins (39.2 mg/kg DW) (León-González et al., 2013; Andrade
et al., 2017). GAE stands for “gallic acid equivalents”. Having a white to colorless fruit, a low
concentration of anthocyanins is to be expected in C. album, as opposed to other colored berries
which have high concentrations of these phenols (Jakobek et al., 2007). The seeds alone have
almost as much phenolic compounds as the entirety of the crowberry fruit. Phenolic compounds
are abundant in C. album leaves (247 mg GAE/100 g leaf weight) and can be found in trace
amounts on the plant flowers (32 mg GAE/100g flower weight) (Brito et al., 2021).The main
phenolic compounds found in C. album fruit are chlorogenic acid, caffeic acid and flavonol
derivatives, mainly myricetin and quercetin (León-González et al., 2013). In the leaves, the
phenols consist mostly of flavonols, such as myricetin and epicatechin (Macedo et al., 2015).
Crowberry phenolics content is found in the Table 7.
Figure 7. Corema album. (A) Crowberry shrub (B) Male flower; (C) Crowberry fruit; (D) Female
flower.
(source: jb.utad.pt; floradegalicia.wordpress.com. photos: A – José Guimarães; B
– João Moleiro;
C – Martin Sirovs; D - Henry David Thoreau)
17
Fatty acids
C. album is an interesting source of healthy lipids. The fatty acids (FA) present in C.
album leaves and fruits are predominantly polyunsaturated (PUFAs), especially in the fruit
seeds, where PUFAs constitute 81.9% of their total fatty acid content (Brito et al., 2021). These
results go in accordance with prior studies by Martin et al. (2019), which revealed that C. album
fruit seeds are rich in unsaturated lipids. Saturated fatty acids (SFAs) are the second most
common fatty acids present in the leaves (31.6%) and fruit pulp (40.4%), and the least common
in the fruit seeds (5.8%). Monounsaturated fatty acids (MUFAs) appear in low quantities in the
leaves (7.77%) and fruit pulp (11.9%) in comparison with the PUFAs and SFAs. The most
common fatty acid in crowberry’s leaves and seeds is alpha-linolenic (leaves: 13.87%; seeds:
50.8%), and in the fruit pulp is eicosapentanoic (30.7%) (Brito et al., 2021). Crowberry fatty
acids content is found in the Table 8.
Reference
[A]
[B]
[C]
Region
Mira, Portugal
Huelva, Spain
Leiria, Portugal
Obs.
White fruit
(X
̄*)
Translucent fruit
(X
̄*)
Fruit
(mg/100 g DW)
Fruit
(mg GAE/100 g DW)
Leaves
(mg GAE/100g FW)
Flowers
(mg GAE/100g FW)
Pulp
Seed
Pulp
Seed
Pulp
Seed
Total phenolic compounds
1517.71
737.31
1342.31
777.31
226.81 ± 22.99
3 ± 1.0
29 ± 1.0
247 ± 21.0
32 ± 2.0
Anthocyanins
-
-
-
-
3.92 ± 0.48
-
-
-
-
Flavonoids
1204.02
546.02
1055.02
531.32
-
-
-
-
-
Flavonols
-
-
-
-
63.83 ± 8.01
-
-
-
-
Tannins
818.33
614.03
732.03
577.03
-
-
-
-
-
Ortho-diphenols
18.81
15.71
17.91
14.81
-
-
-
-
-
[A] Andrade et al., 2017; [B] León-González et al., 2013; [C] Brito et al., 2021.
* Average of three different extraction methods: ethanol:water (50:50, v:v); acetone:water (60:40, v:v); methanol (100%).
1 Gallic acid equivalent (mg GAE/100 g)
2 Quercetin equivalent (mg QE/100 g)
3 mg/100 g
Fatty acid (%)
Fruit pulp
Fruit seeds
Leaves
Polyunsaturated FA
47.0 ± 2.0
81.9 ± 1.1
55.4 ± 1.1
Monounsaturated FA
11.9 ± 1.4
12.3 ± 0.3
7.8 ± 0.3
Saturated FA
40.4 ± 1.3
5.8 ± 0.9
31.6 ± 1.1
Table 7. Phenolic contents in C. album fruits, leaves and flowers.
Table 8. Fatty acid percentage content in C. album fruit and leaves.
18
Terpenoids and alkaloids
Martin et al. (2019) reports the presence of various undetermined terpenoid compounds
in C. album berries using FTIR and Raman analysis. These terpenoids are prevalent in the
cuticular wax of the external fruit skin. Limited research has been published regarding the
description of alkaloids and other nitrogen-containing compounds in C. album. Ferreira (2018)
reports the presence of trigonelline in crowberry leaves (10.2 μg/100 mg) and fruits (2.5 μg/100
mg). This alkaloid is effective in the treatment of diabetes-associated neuropathy (Zhou et al.,
2012) and various types of cancer (Hirakawa et al., 2005; Liao et al., 2015).
Minerals
The mineral content of the crowberry berry is generally higher than more typical berries,
such as blackberries, raspberries and strawberries (Baby et al., 2018). The fruit is a good source
of various minerals per edible weight (EW) such as potassium (230 mg/100 g EW), calcium
(91.2 mg/100 g EW), iron (10.0 mg/100 g EW) and zinc (1.8 mg/100 g EW) (Brito et al., 2021).
C. album fruit also has an high content in copper (2.3 mg/100 g of EW), which is a concerning
factor to human consumption as copper’s negative effects in human cells are well documented
(Agarwal et al., 1989; Pablo Rodriguez et al., 2002; Ahamed et al., 2010). Crowberry mineral
content is found in the Table 9.
1.7. Trichomes
Definition and functions
Trichomes are defined as unicellular or multicellular appendages which extend outwards
from the surface of plant organs and originate exclusively from the epidermal cells (Johnson,
1975; Werker, 2000). Although to different extents, all major terrestrial plants possess
Edible weight (mg/100 g)
Dry weight (mg/100 g)
Calcium (Ca)
91.2 ± 36.1
580 ± 230
Iron (Fe)
10.0 ± 4.8
63.5 ± 30.6
Copper (Cu)
2.3 ± 0.8
14.9 ± 5.0
Zinc (Zn)
1.8 ± 0.6
11.5 ± 3.8
Manganese (Mn)
0.9 ± 0.6
5.8 ± 4.1
Nickel (Ni)
0.4 ± 0.1
2.2 ± 0.3
Strontium (Sr)
1.0 ± 0.4
6.0 ± 2.8
Rubidium (Rb)
0.5 ± 0.3
3.2 ± 1.9
Potassium (K)
230 ± 10
2700 ± 100
Table 9. Mineral content in C. album fruits.
19
trichomes (Johnson, 1975) and the layer of trichomes as a whole in a given plant organ is
referred as the indumentum (Karabourniotis et al., 2020). Trichomes can be found anywhere on
the plant, be it in the stems and leaves or the flowers and fruits. They can also grow on the
subterranean parts of the plant, consisting predominantly of roots hairs, formed by specialized
cells called trichoblasts (Bibikova & Gilroy, 2002).
Trichomes are vital to the plant development, being a primary barrier against hazards such
as herbivores, pathogens and UV irradiation (Xiao et al., 2016) by conferring the plant
epidermis robust structural features and chemical protection (LoPresti, 2016). Various studies
notice that plants produce more trichomes in newer leaves after herbivory damage (Agrawal,
1999; Björkman et al., 2008; Dalin et al., 2008), and trichome secretions compounds may help
to deter pathogen infections (Karamanoli et al., 2005; Kliebenstein et al., 2005; Shepherd &
Wagner, 2007). In contrast, the density of trichomes seems to be related to higher fungal
infection success, as a more dense coat of trichomes provides easier adhesion sites to fungal
cysts and higher humidity levels (Calo et al., 2006; Łaźniewska et al., 2012; Wang et al., 2020).
The production of resinous secretions have been proven to regulate temperature and
transpiration in evergreen xerophytes (Dell & McComb, 1979), and indirectly affect
photosynthesis by eliminating potentially toxic waste in halophytes (Paulino et al., 2020).
Trichomes can also guide and attract pollinators through their secretions (Wagner, 1991). Given
the extensive diversity of trichomes their objective classification is not always easy (Werker,
2000). The major distinction made among trichomes is whether they are glandular or non-
glandular (Wang et al., 2021; Werker, 2000).
Glandular trichomes
Glandular trichomes can be unicellular or multicellular (predominantly the latter) and are
defined as secretory structures that can synthetize and store exudates in specialized secretory
cells, rich in secondary metabolites such as terpenoids, phenols (Karabourniotis et al., 2020;
Wagner et al., 2004; Wagner, 1991) and, to a lesser extent, alkaloids (Laue et al., 2000; Dai et
al., 2010). Being capable of producing secretions rich in secondary metabolites, these glandular
trichomes play a direct role in the protection against herbivory and pathogen infections, water
loss regulation and pollinators attraction, based on the compounds present in the trichomes
secretions (Peter & Shanower, 1998; Dai et al., 2010).
20
Non-glandular trichomes
Non-glandular trichomes are the trichomes whose main function is not the synthesis and
secretion of exudates (Karabourniotis et al., 2020), rather acting as protective physical barriers
(Hanley et al., 2007; Liu et al., 2017a). Non-glandular trichomes are more varied in terms of
their morphology, size and density compared to glandular trichomes, being highly variable
among plant species and within an individual plant itself (Dalin et al., 2008; Werker, 2000).
Non-glandular trichomes can be unicellular and multicellular, branched or unbranched, and can
range from small, soft hairs to stiff thorns and hooks. They can be described based on their
general morphology as needle-like, hook-like, or antenna-like, and based on their shape and
appendage number as simple, peltate, or stellate, among other morphological distinctions (Dalin
et al., 2008; Liu et al., 2017a). Usually more than one type of non-glandular trichomes are
present at the surface of the leaves (Werker, 2000). Although non-glandular trichomes lack a
secretory mechanism, they are reportedly able to store phenolic compounds in vast quantities,
especially during their early stages of development (El-Negoumy et al., 1986; Tattini et al.,
2007; Karabourniotis et al., 2020).
1.8. Objective
The present study objective is to test and assess the anti-oomycete activity of two different
plant species leaf ethanolic extracts fractions against P. cinnamomi. The species studied were
the tree Solanum betaceum, native to the Andes region in South America, and the shrub Corema
album, endemic to the Atlantic coast of the Iberian Peninsula.
21
2. MATERIALS AND METHODS
2.1. Plant material
Material collection locations
The plant material used in this study were the leaves of tamarillo trees and crowberry
bushes. Both leaf types were collected by hand. S. betaceum leaves were collected from twelve
different trees growing at the Botanic Garden of the University of Coimbra (40º12’22”N,
8º25’31”W, altitude 68m) whereas C. album leaves were collected from several bushes at
Quiaios beach (40º13’27”N, 8º53’22”W, altitude 8m). Both collection locations can be seen in
Fig. 8. After collection, both the tamarillo and crowberry leaves were stored in airtight plastic
bags at -40 ºC and later freeze-dried.
Figure 8. A - Tamarillo trees location, represented by the yellow diamond. (1) Botanical Garden
of UC; (2) Departamento de Ciências da Vida of UC.
B – Portuguese-Crowberry bushes location, represented by the yellow area. (3) Quiaios beach.
(source: google.com/earth)
22
Material processing
The leaves were freeze-dried and milled into a fine powder (≤ 1 mm granulometry) using
a blender and a coffee grinder (Qilive Q.5684 and Qilive 870873, respectively), totaling a
weight of 208.88 g of milled tamarillo material and 521.23 g of milled crowberry material, that
accounts for 24.4 % and 38.6 % of the original tamarillo and crowberry material fresh weight,
respectively. The milled material was stored in plastic bags at room temperature (Fig. 9).
2.2. P. cinnamomi cultures and growth medium
P. cinnamomi was freshly propagated from pre-established in vitro cultures (courtesy of
João Martins) and collected as needed throughout the study. The oomycete was propagated in
plastic Petri dishes with Potato Dextrose Agar (PDA) medium (OXOID, LabMal) that consist
of a mixture of potato extract, glucose and agar. P. cinnamomi was left to grow in the dark at
room temperature.
2.3. Plant extract preparation
Compound extraction using ethanol
Both tamarillo material (TAM) and crowberry material (CAM) were weighed in a 2 L
erlenmeyer flask and mixed with ethanol 70 %. 76.7 g of milled TAM material and 100.3 g of
milled CAM material were mixed with 1L of ethanol (Fig. 10A). The mixtures were then stirred
overnight (O/N) on a magnetic stirrer at 110 rpm. On the next day, the mixture supernatants
were decanted via vacuum filtration (VWR VP-86, AVANTOR) (Fig. 10B). To maximize the
Figure 9. Plant milled material. (A) S. betaceum; (B) C. album.
23
compound extraction, the three aforementioned steps were repeated two more times, resulting
in three total ethanol extractions for each plant material. The ethanol extract was stored in Schott
bottles (Fig. 10C) and the solid biomass residue was stored at 4 ºC.
Liquid phase and solid biomass - Ethanol and chloroform extract preparation
To evaporate the ethanol from the extract solution, a rotary evaporator (BÜCHI R-200)
was used with a 40º C heating bath (BÜCHI B-490) (Fig. 11A). The extract evaporation was
done 175 mL at a time in a 500 mL round flask. Due to the substantial volume of ethanol extract
obtained from the prior extraction, the ethanol evaporation was proven to be cumbersome and
time consuming. Both TAM and CAM ethanol evaporated extracts differ greatly from the
ethanol extracts before evaporation, having a cloudy appearance due to the suspension of non-
water-soluble compounds (Fig. 11B). After evaporation, the evaporated ethanol extracts were
freeze-dried until constant weight and labeled 0-EtOH.
Figure 11. Evaporation of the EtOH extracts. (A) Rotary evaporator
apparatus; (B) CAM ethanol extract appearance after evaporation.
Figure 10. Preparation of the ethanol extracts. (A) Mix of milled plant material with
ethanol; (B) Vacuum filtration; (C) The final TAM and CAM ethanol extracts.
24
The initial solid biomasses were subject to an extraction using chloroform only.
Chloroform was added to the biomass and left O/N at around 180 rpm. On the next day the
chloroform was separated via gravity filtration and the solid residue was stored at 4 ºC. Finally,
the chloroform extract was freeze-dried until constant weight and labeled CHL-2.
In the end we were left with two distinct fractions of each TAM and CAM extracts, the
solid biomass residue chloroform extract (labeled CHL-2) and the solid ethanol extract (labeled
0-EtOH). The freeze-dried TAM and CAM ethanol extracts were stored in falcon tubes and the
solid biomass fractions were stored in an airtight bag in the dark at room temperature.
Liquid phase - Fractionation using Liquid Biphasic Systems (LBS)
To prepare the next extracts we resorted to fractionation of the TAM and CAM ethanol
extracts using LBS. The solvents used were n-hexane, chloroform, ethyl acetate and n-butanol,
in this order. The predicted compounds removed by each solvent are shown in the Table 10
below, along each solvent density compared to water.
Solvent
Chemical formula
Density (kg/m3)
Compounds removed
n-Hexane
CH3(CH2)CH3
655.0
Non-polar compounds
Chloroform
CHCl3
1476.8
Pigments
Ethyl Acetate
C4H8O2
902.0
Biological active compounds
n-Butanol
C4H9OH
810.0
Other hydrocarbons
Water
H2O
997.0
-
8 g (TAM) and 10 g (CAM) of freeze-dried ethanol extract (labeled 0-EtOH) were
weighed into 250 mL Schott bottles. 100 mL deionized water was then added to each extract
and stirred vigorously to solubilize/suspend the extract as much as possible. (1) 100 mL of n-
hexane was then added and the (2) solution was stirred for roughly 30 min at >200 rpm to avoid
the formation of layers and ensure an homogenous mixing. (3) The stirred mixtures were then
transferred to a separation funnel and left to rest until separate layers were formed. (Fig. 12) (4)
Both layers were collected into separate bottles, the organic phase (labeled 1-HEX fraction)
into a 100 mL amber bottle and the aqueous phase into a Schott bottle. Because water is denser
than n-hexane (see Table 10 above), the bottom layer will be the aqueous phase, and the top
layer the organic phase. This will be true for the remaining separations except during the
chloroform separation, due to it being denser than water, thus creating an inverted layer order.
Table 10. Solvents used in LBS and predicted compounds removed in each fraction.
Table x. Solvents used in LBS and predicted compounds removed in each fraction.
25
To prepare the next fractions, we added each solvent in succession to the previously collected
aqueous phase, repeating the separation process through steps 1-4 changing the solvent added
in each subsequent separation. All the aforementioned procedures were repeated for both our
freeze-dried TAM and CAM ethanol extracts. In the end, we are left with four different fractions
(labeled 1-HEX, 2-CHL, 3-ETA and 4-BUT) and a remainder aqueous solution (labeled 5-
AQU) of each of our TAM and CAM initial ethanol extracts.
The solvents were then evaporated from the extracts using the rotary evaporator at 40 ºC.
Due to the high boiling point of n-butanol, its fraction was mixed evenly with water to produce
an azeotropic mixture and lower its boiling point before evaporation. The final evaporated
fractions were then reconstituted with acetone and water and left to rest up to a week in the
hotte, to evaporate the acetone from solution at room temperature. Finally, all fractions were
freeze-dried until constant weight and stored in amber flasks. The lyophilized aqueous solutions
(5-AQU) were stored in falcon tubes. The entire extract preparation steps are schematized in
the Fig. 13.
Figure 12. Liquid biphasic system (LBS) fraction separation. (A) Beginning
of the separation (TAM); (B) Fractions fully separated (CAM).
26
Figure 13. Extract preparation step-by-step scheme.
27
2.4. FTIR –Fourier-Transform Infrared Spectroscopy
The chemical characterization of all the TAM and CAM freeze-dried fractions was done
via Fourier-Transform Infrared Spectroscopy (FTIR), using a FTIR spectrometer (VERTEX
70, BRUKER) in junction with an attenuated total reflectance attachment (ATR) (Fig. 14).
FTIR analysis is an effective technique for the chemical analysis of a given sample (Baker et
al., 2014), being a fast, inexpensive and non-destructive procedure (Durak & Depciuch, 2020).
This technique consists in the collection and reading of spectral data over a wide spectral range
to identify the functional chemical groups present in the sampled material (Talari et al., 2017).
These functional groups absorb different wavelengths depending on their type and chemical
bond, that can be detected via spectroscopy and analyzed in the resulting spectra graph.
Although non-compound specific, the identification of the samples molecular functional groups
via FTIR is helpful to narrow the identification of the molecules present in the given sample.
FTIR data analysis and tentative attributions
To analyze each fraction, a small portion of dried material was placed above the
spectrometer laser and the ATR was closed to tightly compress the material and ensure a
reliable reading. The fraction samples were analyzed without further preparation and the spectra
data was recorded in the range of 4000 cm-1 to 400 cm-1. The absorbance spectra were treated
with H2O compensation and CO2 compensation (atmospheric compensation) and converted to
text files in OPUS (v. 7.5; BRUKER OPTIK), imported into MATLAB (v. R2020b;
MATHWORKS, Natick, MA, USA), baseline adjusted using Automatic Weighted Least
Squares (algorithm dim: 2 across columns; polynomial order: 2), and then vector normalized
(Norm: Returns a vector of unit length; length=1). The absorbance spectra were finally cropped
to fingerprint region of 1800 cm-1 to 800 cm-1.
Figure 14. FTIR spectrometer with ATR attachment.
28
2.5. Multiwell Activity Assays
Treatment growth and test model
The effects of the extract fractions on P. cinnamomi growth were tested via multiwell
activity assays (MAA) using 24-well multiwell plates with 16 mm diameter wells (COSTAR,
Corning Inc.), in order to determine each fraction minimum inhibitory concentration (MIC) and
minimum lethal concentration (MLC). Many different fraction concentrations were tested in a
total of six assays, testing three replicates per concentration. The oomycete growth medium
used was PDB, and the fractions were prepared with water. The extract concentrations tested
were honed in each subsequent assay, according to the results obtained in the previous ones.
The first assay was tentatively conducted at concentrations of 10, 5, 2.5 and 1.25 mg/mL for
each fraction but, due to the chemical properties of the different compounds removed in each
fraction, not all fractions could successfully solubilize at this concentration, being diluted at
higher ratios accordingly. As the extract and PDB medium are mixed 1:1 in the wells, both were
priorly prepared at 2x concentration. Each treatment fraction plus PDB solution was mixed
outside of the wells and then equally divided between three wells to conduct the three replicates
per treatment.
As previously noted, not all of the fractions are equally soluble in water, making it
impossible to test some of the extract fractions at high concentrations without the addition of
DMSO, such as the n-hexane, chloroform and ethyl acetate fractions, which were tested at much
lower concentrations when compared to the maximum tested concentrations of ethanol, n-
butanol and aqueous fractions which are highly soluble in water. The solid residue chloroform
fraction (2-CHL) was extremely hard to solubilize in water and was deemed not worthy of
further testing in the present study. As expected, the addition of DMSO affected the oomycete
growth, as this solvent disruptive effect in biological cells is well documented, even at
exceedingly low concentrations (Rammler & Zaffaroni, 1967; Jelke & Oertel, 1990; Galvao et
al., 2014). Additionally, some other tests were conducted without PDB medium, such as extract
only tests and extract plus DMSO.
Control treatments were conducted solely with PDB medium or water. A negative control
assay (C-) was prepared using the commercial fungicide Aliette Flash (BAYER), where five
concentrations were tested: 10, 5, 2.5, 1.25 and 0.75 mg/mL. Aliette Flash active compound
consists of phosphonic acid or fosetyl-Al (Bayer, 2017), a compound belonging into the
phosphite category of compounds known to effectively inhibit P. cinnamomi growth. This
fungicide was the only substance to be tested in negative control treatments, as no purified
29
phosphite compounds were available to test. A positive control (C+) was conducted with PDB
1x medium only. The PDB medium was initially sterilized via autoclaving, and all the extracts,
water and fungicide were filtered during the assays to prevent contamination. The extracts,
water and fungicide were filtered through 0.22 μm polyethersulfone (PES) syringe filters.
Finally, the oomycete was left to grow for 7 days in the dark at room temperature. The MAA
test model is schematized in the Fig. 15 below.
Post treatment growth (PTG)
After the seven days of MAA treatment, several selected P. cinnamomi material (mainly
material which failed to grow at a given fraction concentration) was transferred to regular petri
dishes with PDA medium to assess the oomycete post treatment growth (PTG test). This
additional treatment let us detect if a given concentration corresponds to a lethal concentration
or an inhibitory concentration. If P. cinnamomi fails to grow at a given MAA tested
concentration but subsequentially grow in PTG, the tested concentration is determined to be an
inhibitory concentration. If P. cinnamomi fails to grow in the MAA and in PTG, the tested
concentration is determined to be a lethal concentration. The material transfer was done by
suctioning the growth medium mixture out of the selected treatment well, rinsing the leftover
oomycete material with distilled water two to three times using a pipette, and transferring the
material to a separate petri dish with PDA medium only using a regular tongs. The material was
left to grow in the dark at room temperature for a minimum of five days. The PTG test model
is schematized in the Fig. 16.
Figure 15. Multiwell activity assays (MAA) test model. (1-4) Different
tested concentrations (from highest to lowest); (A, B)
Different extracts.
30
2.6. Extract Fraction Activity Assays
Assay test model
Only three extract fractions from each plant species leaves remained leftover from the
previous multiwell MAA, the ethanol extract, the n-butanol fraction and the aqueous fraction.
From these remaining material, additional activity assays were conducted with the n-butanol
and aqueous fractions to precisely measure their influence on P. cinnamomi hyphal growth area
(HGA). The ethanol extract was very difficult to solubilize in water at the desired
concentrations, and for consistency’s sake it was not tested in this assay. The extract fractions
activity assays (EFAA) were performed in plastic petri dishes with 9 cm diameter, with the
same PDA medium used in the oomycete propagation. Four concentrations of each species n-
butanol and aqueous fractions were tested: 10 mg/mL (1), 5 mg/mL (2), 2.5 mg/mL (3) and
1.25 mg/mL (4). As the fractions are mixed with PDA medium at 1:1 ratio in the petri dishes,
the fractions and PDA medium concentration were previously prepared at 2x the desired
concentration. Concentrations 2, 3 and 4 were prepared by sequential water dilutions, starting
from the original concentration.
15 mL of each fraction were mixed with 15 mL of PDA 2x medium in small Erlenmeyer
flasks, distributed evenly in Petri dishes and left to solidify. A total of three replicates per extract
concentration were conducted. Each Petri dish totaled roughly 10 mL of fraction plus PDA.
Three control (C+) replicates were prepared using only PDA 1x medium. As a matter of interest,
three additional C+ replicates were prepared with leftover PDA 2x medium. Finally, small equal
circles of P. cinnamomi were cut and collected from our previous cultures, placed at the center
of the prepared Petri dishes. The water, PDA medium and laboratory material used were
initially sterilized via autoclaving to prevent contamination. The EFAA test model is
schematized in the Fig. 17.
Figure 16. Post treatment growth (PTG) test model.
31
Growth area and statistical analysis
P. cinnamomi was left to grow for 5 days in the dark at room temperature. After treatment,
each individual Petri dish was photographed from top view at a height of 7.5 cm, in the same
position and light conditions with the help of a camera stand and a regular smartphone camera
(Fig. 18). P. cinnamomi mycelium hyphal growth area was measured and analyzed using
ImageJ1 software (NIH) (Hartig, 2013) (Fig. 19) and the results are expressed in cm2. Growth
reduction was calculated from the three replicates in accordance with Royse & Ries (1978),
using the formula: GR = (TG-CG)/CG *100, where GR, TG and CG stand for “growth
reduction”, “treatment growth” and “control growth”, respectively.
The obtained growth area data was subject to one-way analysis of variance (ANOVA)
and post hoc analysis using Tuckey test, both with p = 0.05. Tuckey test p-values were corrected
according to Bonferroni (1936) using the formula α = P/N, where P and N note the “p
significance” and “number of comparisons” between groups, respectively. The statistical
analysis was conducted using the Data Analysis ToolPak add-on of Microsoft Excel in
accordance with Salkind (2015).
Figure 18. Photo capture model. (1) Camera
stand; (2) Camera; (3) Petri dish.
Figure 19. ImageJ1 software interface with a
loaded and processed image.
Figure 17. Extract fraction activity assays (EFAA) test model. (1-4) Different concentrations.
32
2.7. Trichome observation and scanning electron microscopy
Sampled material
A total of four S. betaceum varieties leaf material were sampled for SEM observation:
Two varieties based on the fruit color (red-type and orange-type), and two in vitro varieties (C9
and C12) (courtesy of Dr. Sandra Correia). Red-type and orange-type varieties were sampled
from leaves of the same tamarillo trees from which we previously collected leaves for extract
preparation planted at the Botanical Garden of UC. The two in vitro genotypes were pre-
established seedlings grown in test tubes of unknown age (Fig. 20). A single variety of C. album
leaf material was observed, sampled from plant material collected at Praia de Quiaios.
Scanning electron microscopy
Scanning electron microscopy took place at Instituto Biomédico de Investigação, Luz e
Imagem (IBILI) using a variable pressure scanning electron microscope (Flex SEM 1000,
HITACHI). Samples were prepared by cutting small square segments of leaf blade and several
cross-sections along the midrib, placed on carbon stickers above metallic stubs and observed
without further preparation. The observations were conducted at 10.0 kV in freeze conditions
(-20 °C).
Figure 20. In vitro tamarillo varieties observed in SEM.
(A) C9 variety; (B) C12 variety.
33
3. RESULTS
3.1. Multiwell Activity Assays
The treatments tested in the multiwell activity assays (MAA) can be divided into three
major categories, named here as “standard” treatments (extract fraction plus PDB medium
only), “conditioned” treatments (any treatment conducted using the extract fraction
with/without DMSO and/or PDB medium) and control treatments. It is important to give this
distinction between “standard” and “conditioned” treatments, as in both species the oomycete
growth is significantly influenced by whether or not DMSO and PDB medium are present in
the treatment. All individual MAA results and test models can be found in Appendix 1-6.
3.1.1. Solanum betaceum MAA
Standard treatments
In the standard treatments with S. betaceum fractions, the n-butanol fraction inhibited P.
cinnamomi growth, at concentrations of 18 mg/mL. The oomycete also failed to grow in a
concentration of 3.33 mg/mL with the ethyl acetate fraction in two of the three total replicates
conducted. P. cinnamomi material treated with the ethyl acetate fraction at 3.33 mg/mL and
with the n-butanol fraction at 18 mg/mL were subjected to post-treatment growth tests (PTG).
P. cinnamomi tested with the ethyl acetate fraction at 3.33 mg/mL and the n-butanol fraction at
18 mg/mL successfully grew in PTG tests. All the remaining treatments conducted with S.
betaceum fractions failed to inhibit P. cinnamomi growth at tested concentrations. All results
from the standard treatments with S. betaceum fractions are shown in Table 11 and the
compiled tamarillo fractions MIC/MLC is found in the Table 12.
34
Fraction
C (mg/mL)
Growth
PTG
1-HEX
(n-Hexane)
1.43
+
0.715
+
0.358
+
0.1788
+
2-CHL
(Chloroform)
1
+
0.5
+
0.25
+
0.125
+
3-ETA
(Ethyl acetate)
3.33
+
+
1.665
+
0.833
+
0.416
+
4-BUT
(n-Butanol)
18
-
+
10.95
+
10.90
+
10.85
+
10.80
+
10.75
+
10.50
+
10.25
+
10
+
9
+
5
+
4.5
+
2.5
+
2.25
+
1.25
+
5-AQU
(Aqueous)
18
+
16
+
14
+
12
+
10
+
5
+
2.5
+
1.25
+
0-EtOH
(Ethanol)
18
+
16
+
14
+
12
+
10
+
5
+
2.5
+
1.25
+
+ Growth
- No growth
Leaf
material
Fraction
MIC
(mg/mL)
MLC
(mg/mL)
Tamarillo
(S. betaceum)
Ethanol
> 18
-
n-Hexane
> 1.43
-
Chloroform
> 1
-
Ethyl Acetate
> 3.33
-
n-Butanol
10.95 - 18
> 18
Aqueous
> 18
-
Table 11. MAA and PTG results in standard treatments with S. betaceum fractions.
Table x. MAA and PTG results in standard treatments with S. betaceum fractions.
Table 12. S. betaceum fractions MIC and MLC.
35
Conditioned treatments
In the conditioned tests with S. betaceum extracts, two main types of treatments were tested,
the combination of DMSO plus fraction with PDB medium and DMSO plus fraction without
PDB medium. A third type of treatment was conducted with n-butanol fraction exclusively
between 11-14 mg/mL, without DMSO and PDB. Three of the conducted treatments
successfully inhibited P. cinnamomi growth, DMSO 0.25 % plus n-hexane 1.5 mg/mL without
PDB, DMSO 5 % plus ethyl acetate 6 mg/mL with PDB and DMSO 0.5 % plus ethyl acetate 6
mg/mL without PDB. All the treatments with the standalone n-butanol fraction inhibited P.
cinnamomi growth. Further PTG tests were conducted in oomycete material tested with n-
butanol at 11 and 12 mg/mL, in which the material successfully grew. The results from the
conditioned treatments conducted with S. betaceum fractions are shown in Table 13.
Fraction
C (mg/mL)
Observations
Growth
PTG
1-HEX
(n-Hexane)
3
DMSO 5 %
+
1.5
DMSO 5 %
+
1.5
DMSO 0.25 % (without PDB)
-
0.75
DMSO 5 %
+
0.75
DMSO 0.25 % (without PDB)
+
0.375
DMSO 5 %
+
0.375
DMSO 0.25 % (without PDB)
+
0.1875
DMSO 0.25 % (without PDB)
+
2-CHL
(Chloroform)
2
DMSO 5 %
+
1
DMSO 5 %
+
1
DMSO 0.25 % (without PDB)
+
0.5
DMSO 5 %
+
0.5
DMSO 0.25 % (without PDB)
+
0.25
DMSO 5 %
+
0.25
DMSO 0.25 % (without PDB)
+
0.125
DMSO 0.25 % (without PDB)
+
3-ETA
(Ethyl acetate)
6
DMSO 5 %
-
6
DMSO 0.5 % (without PDB)
-
3
DMSO 5 %
+
1.5
DMSO 5 %
+
0.75
DMSO 5 %
+
4-BUT
(n-Butanol)
14
without PDB
-
13
without PDB
-
12
without PDB
-
+
11
without PDB
-
+
+ Growth
- No growth
Table 13. MAA results in conditioned treatments with S. betaceum fractions.
Table x. MAA results in conditioned treatments with S. betaceum fractions.
36
3.1.2. Corema album MAA
Standard treatments
The ethanol, n-hexane, chloroform and ethyl acetate fractions successfully inhibited P.
cinnamomi growth at some given concentration. Both the n-hexane and the chloroform fractions
inhibited P. cinnamomi growth at a concentration of 1.75 mg/mL. The ethanol and ethyl acetate
fractions inhibited the oomycete growth at 1.25 mg/mL and 0.7 mg/mL, respectively. Oomycete
material treated with the chloroform fraction at 1.25 mg/mL and above, with the ethyl acetate
fraction at 1.25 mg/mL and above (except the 1.5 mg/mL treatment), with the n-butanol fraction
between 10-10.75 mg/mL and with the ethanol extract at 0.7 mg/mL and above, were all subject
to additional PTG tests. Regarding the PTG of material treated with the chloroform fraction,
only the oomycete material priorly treated at 1.25 mg/mL grew. No P. cinnamomi material
treated with the ethyl acetate fraction grew during PTG tests. During PTG tests in material
treated with the n-butanol fraction, only the oomycete material treated at 1.5 mg/mL failed to
grow. In PTG tests conducted with P. cinnamomi material previously treated with the ethanol
extract, material treated between 0.7-1.1 mg/mL failed to grow and only the material treated at
1.25 and 2.5 mg/mL grew. All results from the standard treatments with C. album fractions are
shown in Table 14 and the compiled tamarillo fractions MIC/MLC is found in the Table 15.
37
Fraction
C (mg/mL)
Growth
PTG
1-HEX
(n-Hexane)
4
-
2
-
1.75
-
1.50
+
1.25
+
1
+
0.5
+
2-CHL
(Chloroform)
2.5
-
-
2
-
-
1.75
-
-
1.50
+
-
1.25
+
+
0.625
+
0.313
+
3-ETA
(Ethyl acetate)
2
-
-
1.75
-
-
1.50
-
1.25
-
-
1
+
0.5
+
0.25
+
4-BUT
(n-Butanol)
18
a
16
a
14
a
12
a
10.75
a
+
10.50
a
-
10.25
a
+
10
+
+
5
+
2.5
+
1.25
+
5-AQU
(Aqueous)
18
+
16
+
14
+
12
+
10
+
5
+
2.5
+
1.25
+
0-EtOH
(Ethanol)
2.5
-
+
1.25
-
+
1.1
-
-
1
-
-
0.9
-
-
0.8
-
-
0.7
-
-
0.625
+
0.3125
+
+ Growth
- No growth
a Unknown, indiscernible
Table 14. MAA and PTG results in standard treatments with C. album fractions.
Conflicting results between MAA and PTG tests are presented in red.
Table x. MAA and PTG results in standard treatments with C. album fractions.
Conflicting results between MAA and PTG tests are presented in red.
38
Leaf
material
Fraction
MIC
(mg/mL)
MLC
(mg/mL)
Crowberry
(C. album)
Ethanol
0.625 - 0.7
0.7
n-Hexane
1.75
-
Chloroform
1.5 - 1.75
1.75
Ethyl Acetate
1.00 - 1.25
1.25
n-Butanol
> 10
> 10.75
Aqueous
> 18
-
Conditioned treatments
A single conditioned test was conducted using the n-hexane fraction, at a concentration
of 0.25 mg/mL without PDB, in which P. cinnamomi successfully grew. Several treatments
without PDB were conducted with the n-butanol fraction starting at a concentration of 11
mg/mL. No oomycete growth was observed in any of these conditioned treatments using the n-
butanol fraction. The n-butanol fraction treatments conducted at 11 and 12 mg/mL were subject
to further PTG tests, in which they failed to grow. The results from the conditioned treatments
conducted with C. album fractions are shown in Table 16.
Fraction
C (mg/mL)
Observations
Growth
PTG
1-HEX
0.25
without PDB
+
4-BUT
18
without PDB
-
16
without PDB
-
14
without PDB
-
13
without PDB
-
12
without PDB
-
-
11
without PDB
-
-
+ Growth
- No growth
3.1.3. Aliette Flash and DMSO MAA
The negative control treatments conducted using Aliette Flash successfully inhibited P.
cinnamomi growth, even at the lowest concentration tested of 0.75 mg/mL. No further PTG
tests were conducted in the material tested with Aliette Flash. Additional control tests were
conducted using DMSO only to assess the compound standalone effects on P. cinnamomi
growth. The oomycete is able to grow in DMSO concentrations as high as 2.5 % and fails to
grow at 5 % DMSO concentration, with similar results observed in the PTG tests conducted in
this material. Both treatments results are found in the Table 17.
Table 16. MAA and PTG results in conditioned tests with C. album fractions.
Table x. MAA and PTG results in conditioned tests with C. album fractions.
Table 15. C. album fractions MIC and MLC.
39
Fraction
C
Growth
PTG
Aliette
Flash
10
-
5
-
2.5
-
1.25
-
0.75
-
DMSO
5
-
-
2.5
+
+
2.0
+
+
1.5
+
+
0.5
+
+
+ Growth
- No growth
3.2. Extract Fraction Activity Assays
The final determined treatment growth (TG) is presented alongside each treatment growth
reduction (GR) when compared to control growth (CG). TG is noted in cm2, and GR is noted
both in cm2 and percentage (%). GRmean and its associated standard deviation (StdDev) are
obtained from each treatment three replicates GR in percentage (%). Statistically significant
differences between treatments are noted as different letters in the result graphs. The complete
HFA growth area results can be found in Appendix 7. The full ANOVA and Tuckey test data
from the EFAA can be found in Appendix 8-19.
3.2.1. Control EFAA
P. cinnamomi grew on average 22.8 cm2 in the control treatment with PDA, reaching a
maximum hyphal growth area (HGA) of 23.1 cm2 and a minimum HGA of 22.6 cm2. P.
cinnamomi grew to an average of 16.2 cm2 in PDA 2x control treatments, with a maximum
HGA of 18.8 cm2 and a minimum HGA of 13.8 cm2 (Table 18 and Fig. 21).
Control
CG (cm2)
CGmean (cm2)
StdDev
PDA
22.679
22.813
0.218
22.639
23.121
PDA 2x
16.054
16.198
2.060
18.789
13.750
Table 17. MAA and PTG results in Aliette Flash and DMSO treatments. Aliette Flash
and DMSO concentrations are noted in mg/mL and in percentage (%), respectively.
Table x. MAA and PTG results in DMSO treatments.
Table 18. EFAA control treatments hyphal growth area results.
Table x. EFAA control treatments hyphal growth area results.
40
3.2.2. Solanum betaceum EFAA
S. betaceum n-butanol fraction decreases P. cinnamomi HGA by 7.67 ± 3.39 % in the
1.25 mg/mL treatment, 41.57 ± 8.12 % in the 2.5 mg/mL treatment, 64.35 ± 4.06 % in the 5
mg/mL treatment and 75.90 ± 1.18 % in the 10 mg/mL treatment. Statistical analysis shows
that the oomycete HGA is significantly different between 1.25, 2.5 and 10 mg/mL treatments,
and not significantly different between 2.5-5 mg/mL and 5-10 mg/mL (Fig. 22 and 23).
Figure 21. Control treatments growth after 5 days. (A) PDA; (B) PDA 2x.
Figure 23. S. betaceum n-butanol fraction treatments. A single replicate from the
total three is shown. (A) 10 mg/mL; (B) 5 mg/mL; (C) 2.5 mg/mL; (D) 1.25 mg/mL.
Figure 22. S. betaceum n-butanol fraction GR in percentage.
41
S. betaceum aqueous fraction reduces P. cinnamomi HGA by 8.60 ± 10.35 % in the 1.25
mg/mL treatment, 22.36 ± 2.58 % in the 2.5 mg/mL treatment, 36.53 ± 7.07 % in the 5 mg/mL
treatment and 53.00 ± 3.41 % in the 10 mg/mL treatment. One of the 1.25 mg/mL treatment
replicates shows a HGA increase of 6.60 % was detected when compared to the control. The
statistical analysis shows that there were no significant differences between the HGA of 1.25,
2.5 and 5 mg/mL treatments, as well as between 5-10 mg/mL, but the HGA of the 1.25 and 2.5
mg/mL treatments are significantly different from 10 mg/mL (Fig. 24 and 25).
Figure 25. S. betaceum aqueous fraction treatments. A single replicate from the
total three is shown. (A) 10 mg/mL; (B) 5 mg/mL; (C) 2.5 mg/mL; (D) 1.25 mg/mL.
Figure 24. S. betaceum aqueous fraction GR in percentage.
42
3.2.3. Corema album EFAA
C. album n-butanol fraction decreases P. cinnamomi HGA by 65.81 ± 4.41 % in the 1.25
mg/mL treatment, 81.24 ± 2.16 % in the 2.5 mg/mL treatment, 89.37 ± 1.14 % in the 5 mg/mL
treatment and 100 % in the 10 mg/mL treatment, where it completely inhibits oomycete growth.
There were no statistically significant differences between 1.25-2.5 mg/mL treatments HGA,
but all the remaining treatments are significantly different amongst themselves (Fig. 26 and
27).
Figure 26. C. album n-butanol fraction GR in percentage.
Figure 27. C. album n-butanol fraction treatments. A single replicate from the total
three is shown. (A) 10 mg/mL; (B) 5 mg/mL; (C) 2.5 mg/mL; (D) 1.25 mg/mL.
43
C. album aqueous fraction increases or diminishes P. cinnamomi HGA depending on the
treatment concentration. When compared to the control, the treatment at 1.25 mg/mL increases
HGA by 9.02 ± 2.37 %. P. cinnamomi HGA decreases by 1.49 ± 14.60 % in the 2.5 mg/mL
treatment, 32.94 ± 2.76 % in the 5 mg/mL treatment and 57.89 ± 2.58 % in the 10 mg/mL
treatment. There were no statistically significant differences between 1.25-2.5 mg/mL and 2.5-
5 mg/mL treatments, being the remaining treatments significantly different amongst each other
(Fig. 28 and 29).
Figure 28. C. album aqueous fraction GR in percentage.
Figure 29. C. album aqueous fraction treatments. A single replicate from the total
three is shown. (A) 10 mg/mL; (B) 5 mg/mL; (C) 2.5 mg/mL; (D) 1.25 mg/mL.
44
3.3. FTIR analysis
Only the FTIR spectra obtained from fractions which were able to influence P. cinnamomi
growth during the MAA and the fractions used in the EFAA will be thoroughly analyzed. The
attributions go in accordance with Martin et al. (2019) and Martin et al. (2021). The FTIR
spectra of all the remaining fractions that are not presented can be found in Appendix 20-25.
3.3.1. Solanum betaceum FTIR analysis
n-Butanol fraction spectrum
The n-butanol fraction spectrum presents many narrow and pronounced peaks throughout
the entire fingerprint region. The most prominent peak occurs at 1596 cm-1 (a) which suggests
the presence of polyphenols in the fraction. Next, the peak at 1260 cm-1 (c) could be found with
a shoulder protruding at 1282 cm-1 (b), both corresponding to the presence of bending hydroxyl
(OH) groups, suggesting the presence of polysaccharides such as fructose or the presence of
cutin. Right beside this peak, two more paired strong and narrow peaks could be found at 1178
cm-1 (d) and 1160 cm-1 (e), both associated with the presence of antisymmetric stretching in C-
O-C groups corresponding to ester compounds. Finally, one of the more prominent peaks was
detected at the start threshold of the fingerprint region, at 809 cm-1 (j). This final peak may
indicate the presence of triterpenoid compounds. The peaks at 1115 (f) cm-1, 1067 cm-1 (g) and
1040 cm-1 (h) are also worth noting. These peaks suggest the presence of stretching symmetric
C-O-C, stretching C-O and C-C groups, each corresponding to ester and glycosidic bonds,
likely linked to polysaccharides compounds, respectively (Fig. 30).
Aqueous fraction spectrum
The aqueous fraction FTIR spectrum presents significantly less peaks in comparison with
the previous n-butanol fraction. Nonetheless, the same strong peak appears in the aqueous
fraction, at 1596 cm-1 (a) which notes the presence of polyphenols. The peak at 1040 cm-1 (h)
is very pronounced and corresponds to C-C groups associated with the existence of
polysaccharides in the aqueous fraction. This peak also has two prominent shoulders, one at
1067 cm-1 (g) and one at 989 cm-1 (i), associated with the presence of esters and sucrose,
respectively. Two smaller peaks should also be noted: the first at 1260 cm-1 (c) with a shoulder
at 1282 cm-1 (b) associated with the presence of polysaccharides, which also appeared much
more pronounced in the n-butanol fraction, and the second peak at 809 cm-1 (j) pointing to the
presence of triterpenoids (Fig. 31).
45
Wavenumber
(cm-1)
Reference
Biomass
Feature
Tentative
Assignment
a
1596
1609 (Martin et al., 2019)
Crowberry fruit (outer skin)
ν(C═C)ring
Polyphenols
b
1282
1277 (Martin et al., 2019)
Crowberry fruit (outer skin)
δ(OH)
Fructose/
Polysaccharides; cutin
c
1260
1262 (Martin et al., 2021)
Tamarillo fruit (peel)
δ(O-H)
-
d
1178
1175 (Martin et al., 2021)
Tamarillo fruit (outer skin)
ν asym(C-O-C)ester
-
e
1160
1160 (Martin et al., 2021)
Tamarillo fruit (inner skin)
ν asym(C-O-C)ester
-
f
1115
1111 (Martin et al., 2021)
Tamarillo fruit (inner skin)
ν sym(C-O-C)ester
-
g
1067
1068 (Martin et al., 2021)
Tamarillo fruit (peel)
ν(C-O-C)glycosidic
-
h
1040
1038 (Martin et al., 2021)
Tamarillo fruit (seed and inner skin)
ν(C-O) /ν(C-C)
Polysaccharides,
pectins
i
989
995 (Martin et al., 2021)
Tamarillo fruit (peel)
ν(CO) / ν(CC)ring
Sucrose
j
809
808 (Martin et al., 2019)
Crowberry fruit (outer skin)
-
Triterpenoids
Abbreviations: sym symmetric; asym antisymmetric; ν stretching; δ bending; ρ plane; oρ out-of-plane.
Table 19. FTIR analysis and peak values of different functional groups obtained from the n-butanol and
aqueous leaf fractions of S. betaceum. Assignments based on Martin et al. (2019) and Martin et al. (2021).
46
Figure 30. FTIR absorbance spectrum of S. betaceum n-butanol fraction.
Figure 31. FTIR absorbance spectrum of S. betaceum aqueous fraction.
47
3.3.2. Corema album FTIR analysis
Ethanol extract spectrum
The ethanol extract spectrum presents two notable peak clusters around the 1600 cm-1 and
1000 cm-1 regions. In the first region there are two peaks at 1632 cm-1 (a) and 1603 cm-1 (b).
The peak at 1632 cm-1 corresponds to antisymmetric stretching bonds in COO- groups,
suggesting the presence of acids in the fraction. It may also correspond to the presence of water
molecules. The 1603 cm-1 region does not have any assignments in Martin et al. (2019) and
Martin et al. (2021). Nonetheless, previous works by Ahmad et al. (2016) in H. bacciferum
flowers assign the peak region to stretches in C-C ring bonds, suggesting the presence of
aromatic compounds. In the second peak cluster of the ethanol extract spectrum there are four
main peaks: three stronger peaks at 1065 cm-1 (h), 1041 cm-1 (i) and 1030 cm-1 (j) associated
with stretching glycosidic bonds in ester groups (C-O-C), and a weaker peak at 986 cm-1 (k)
that points the existence of stretching linear chains of carbons (C-C) that may denote the
presence of polysaccharides (Fig. 32).
n-Hexane fraction spectrum
The n-hexane fraction spectrum has many narrow peaks with accentuated dips in between
them. Two noteworthy peaks can be observed at 1632 cm-1 (a) and 1159 cm-1 (e). The first peak
can also be found in the ethanol extract spectrum and corresponds to antisymmetric stretching
bonds in COO- groups, and the second peak is associated with stretching CO and CH bonds,
noting the presence of phenolic compounds and polysaccharides such as pectins (Fig. 33).
Chloroform fraction spectrum
The chloroform fraction spectrum depicts two strong peak clusters and many narrow and
prominent peaks. The first cluster hold two peaks at 1632 cm-1 (a) and 1603 cm-1 (b) akin to the
ethanol extract spectrum. The peak at 1632 cm-1 corresponds to COO- groups and the peak at
1603 cm-1 corresponds to the presence of aromatic compounds. The second peak cluster has
three peaks: 1208 cm-1 (d) which suggests the existence of bending CCH bonds associated with
lipid compounds, 1159 cm-1 (h) representing the presence of stretching bonds in CO and CH
groups which suggest the presence of pectins and phenolic compounds, and 1140 cm-1 (f)
associated with stretching glycosidic bonds in COC groups. Two narrow peaks should be
pointed out, one at 1448 cm-1 (c) that may be associated with the presence of lipids and one at
818 cm-1 (l) noting the presence of triterpenoids (Fig. 34).
48
Ethyl acetate fraction spectrum
Three very strong peaks could be seen in the ethyl acetate fraction spectrum. As in
aforementioned fractions spectra, two of the major ethyl acetate fraction peaks are found at
1632 cm-1 (a) and 1603 cm-1 (b), which assignments were already extensively mentioned in the
previous fractions, and the third major peak is found at 1140 cm-1 (f) that is related to the
presence of phenolic compounds and polysaccharides like pectins. Several relatively strong
peak clusters can also be observed around peaks at 1208 cm-1 (d) and 1065 cm-1 (h), which
suggest the presence of lipids and glycosidic bonds in ester groups (COC). A fourth major peak
is found at 818 cm-1 (l) pointing to the existence triterpenoid compounds (Fig. 35).
n-Butanol fraction spectrum
The n-butanol fraction spectrum showed a narrow lone peak at 1603 cm-1 (b)
correspondent to the presence of C-C ring related to aromatic compounds. An even stronger
three peak cluster can be observed in the lower fingerprint threshold, consisting of the peaks at
1041 cm-1 (i), 1030 cm-1 (j) and 986 cm-1 (k), all associated with the presence of
polysaccharides. A protruding shoulder at 1116 cm-1 (g) also suggests the presence of
polysaccharides. The peak a 818 cm-1 (l) noting the existence of triterpenoids is also somewhat
strong, but smaller than in previous fractions (Fig. 36).
Aqueous fraction spectrum
The aqueous fraction spectrum greatly differs from all the other fraction spectra. The
peaks are wide and smooth, with two noteworthy peaks at 1603 cm-1 (b) and 1030 cm-1 (j),
which point the presence of aromatic compounds and the presence of stretching bonds in linear
chains of carbons (C-C), respectively. One strong shoulder can be observed adjacent to the later
peak at 986 cm-1 (k) which corresponds to stretching CO bonds depicting to the presence of
polysaccharides (Fig. 37).
49
* Due to the wavenumbers relevance in the fractions spectra, and the assignments not being found in the
previous studies of Martin et al. (2019) and Martin et al. (2021), from where the present study tentative
assignments are drawn from, this exceptional attributions were taken in accordance with previous studies
in H. bacciferum flowers by Ahmad et al. (2016) and P. angulata stems by Leite et al. (2018).
Wavenumber
(cm-1)
Reference
Biomass
Feature
Tentative assignment
a
1632
1632 (Martin et al., 2021)
Tamarillo fruit (peel)
ν asym (COO-), H2O
- ; water
b
1603
1600 (Ahmad et al., 2016)
Heliotropium bacciferum flower
C-C stretch in ring
Aromatic compounds
c
1448
1444 (Martin et al., 2019)
Crowberry fruit (inner skin)
δ(CH2) /
δ(CH3)glucosydic
Lipids
d
1208
1214 (Martin et al., 2019)
Crowberry fruit (outer skin)
δ(CCH)
Lipids
e
1159
1153 (Martin et al., 2021)
Tamarillo fruit (skin)
ν(CO) / ν(CH)
Pectins
Phenolic compounds
f
1140
1145 (Leite et al., 2018)
Physalis angulata stem
-
Pectins
g
1116
1111 (Martin et al., 2019)
Crowberry fruit (inner skin)
ν(COC)glycosidic
-
h
1065
1065 (Martin et al., 2021)
Tamarillo fruit (skin)
ν(C-O-C)glycosidic
-
i
1041
1047 (Martin et al., 2019)
Crowberry fruit (outer skin)
ν(COC)glycosidic
-
j
1030
1032 (Martin et al., 2019)
Crowberry fruit (inner skin)
ν(C─C)linear chains
Polysaccharides
k
986
976 (Martin et al., 2019)
Crowberry fruit (outer skin)
ν(CO)
Polysaccharides
l
818
808 (Martin et al., 2019)
Crowberry fruit (outer skin)
-
Triterpenoids
Abbreviations: sym symmetric; asym antisymmetric; ν stretching; δ bending; ρ plane; τ twisting; ω wagging.
Table 20. FTIR analysis and peak values of different functional groups obtained from ethanol, n-hexane,
chloroform, ethyl acetate , n-butanol and aqueous leaf fractions of C. album.
*
*
50
Figure 32. FTIR absorbance spectrum of C. album ethanol extract.
Figure 33. FTIR absorbance spectrum of C. album n-hexane fraction.
51
Figure 34. FTIR absorbance spectrum of C. album chloroform fraction.
Figure 35. FTIR absorbance spectrum of C. album ethyl acetate fraction.
52
Figure 36. FTIR absorbance spectrum of S. C. album n-butanol fraction.
Figure 37. FTIR absorbance spectrum of C. album aqueous fraction.
53
3.4. Trichome identification
3.4.1. Solanum betaceum trichomes
Three types of trichomes can be observed in red-type and orange-type of tamarillo leaves,
identified in this study as type A1, A2 and B: Type A1 are uniseriate capitate long glandular
trichomes with four coplanar secretory cells at their tip (e.g.: Fig. 39B), type A2 are short
capitate glandular trichomes, also with four coplanar secretory cells (e.g.: Fig. 38A) and type
B are very long uniseriate apiculate non-glandular trichomes (e.g.: Fig. 38C). A third type of
glandular trichome was observed in the C9 and C12 in vitro S. betaceum varieties, identified
here as type A3 This type is a very long uniseriate capitate glandular trichome, possibly with a
single secretory cell at its tip (e.g.: Fig. 41A).. In all the tamarillo varieties observed, the most
numerous trichome is the non-glandular type B, and the indumentum is significantly denser in
the abaxial leaf surface and along the leaves midrib and veins. C9 and C12 varieties have much
less trichomes than the red and orange-type varieties.
Figure 38. Red-type S. betaceum leaf trichomes
observation in SEM. (A) Transversal cut;
(B)
Adaxial leaf surface; (C, D)
Abaxial leaf surface.
Figure 39. Orange-type S. betaceum leaf trichomes
observation in SEM. (A-D) Transversal cut.
54
3.4.2. Corema album trichomes
Three type of trichome can be observed in C. album, identified in present study as type
C1, C2 and D: Type C1 are long uniseriate capitate glandular trichomes (e.g.: Fig. 42F), type
C2 are short capitate glandular trichomes (Fig. 42G), and type D are long uniseriate non-
glandular trichome which covers the vast majority of the abaxial leaf surface (e.g.: Fig 42E).
Type C1 and C2 are very sparce, especially type C2, which can only be observed once in all
SEM captured observations. C. album adaxial leaf surface is completely glabrous (Fig. 42D).
Figure 40. C9 S. betaceum in vitro variety leaf
trichomes observation in SEM. (A)
Transversal
cut; (B) Adaxial leaf surface; (C, D)
Abaxial leaf
surface.
Figure 41. C12 S. betaceum in vitro variety leaf
trichomes observation in SEM. (A, B)
Transversal
cut; (C, D) Adaxial leaf surface.
Figure 42. C. album leaf trichomes observation in SEM. (A-C) Transversal cut; (D) Adaxial leaf
surface; (E-H) Abaxial leaf surface.
55
4. DISCUSSION
P. cinnamomi is a hazardous phytopathogen responsible for several of plant diseases
across the globe and have a broad affinity to many plants hosts. The consequences onset by
these diseases can be severely detrimental to the habitats were the pathogen hosts inhabit, as
plants hold a fundamental role at the bottom of food chains (Power, 1992) and can provide
shelter to many other coexistent species (Bossenbroek et al., 1977; Fukui, 2001). The oomycete
can also infect plenty of plant crops and garden species, which have an important economic
value. Given the lack of favorable and effective measures to control the propagation of P.
cinnamomi, the search and study of definitive and environmentally friendly solutions is of
utmost importance. A possible solution to this matter is the use of plant extracts as a cheap and
low residual alternative to the use of potentially harmful pesticides (Hossain et al., 2017;
Laxmishree & Nandita, 2017). In the present study, S. betaceum and C. album fractioned leaf
extracts were assessed in vitro conditions to test anti-oomycete activity in P. cinnamomi growth.
To the best of my knowledge, this is the first report in the anti-oomycete activity of S. betaceum
and C. album leaf extracts against P. cinnamomi.
Results analysis - S. betaceum fractions
The n-butanol fraction was the only S. betaceum fraction which showed some kind of
significant inhibitory activity against P. cinnamomi during the MAA. The n-butanol fraction
inhibited P. cinnamomi growth at 18 mg/mL in standard treatments and at 11 mg/mL and above
without PDB in conditioned treatments. Despite these results, no clear conclusions can be
drawn regarding the MIC/MLC of the n-butanol fraction other than it is between 10.95-18
mg/mL, as the gap between this tested concentrations is very high, and the absence of growth
medium highly compromises the oomycete growth. Nonetheless, the n-butanol fraction held
promising results during EFAA growth reduction tests, causing a reduction of 41.57 ± 8.12 %
in P. cinnamomi hyphal growth area (HGA) at concentrations as low as 2.5 mg/mL, and
significantly reduces P. cinnamomi HGA by 75.90 ± 1.18 % at 10 mg/mL. The polyphenols
and terpenoid compounds present in the n-butanol fraction are likely involved in the anti-
oomycete activity observed, as the anti-oomycete activity of both compounds classes are well
documented (e.g.: Shim et al., 2009; Damian Badillo et al., 2010; Madrid et al., 2015;
Montenegro et al., 2019). The presence of polysaccharides may also play a role in the
bioactivity of the n-butanol fraction. Although few, some studies have already been published
56
which report the direct antimicrobial activity of polysaccharides (Paris et al., 2019). Like in the
MAA, the n-butanol fraction and PDA mixture also formed a precipitate during the EFAA, but
in this treatments the precipitate starts forming at a concentration of 5 mg/mL, approximately
at half the concentration at which the precipitate formed during MAA. This is likely due to the
medium used being PDA and not PDB. The S. betaceum ethyl acetate fraction inhibited P.
cinnamomi growth in two of the three replicates conducted at 3.33 mg/mL during the MAA
standard treatments. Despite the inhibition observed in two of the replicates at this
concentration, we could not consider the 3.33 mg/mL as the MIC of the ethyl acetate fraction,
as one of the replicates did grow. The fraction plus PDB medium solution used in the treatments
was primarily mixed outside of the wells and then equally divided between the multiwells,
which means that there were no possible treatment differences between each replicate and being
plausible that the remaining two replicates failed to grow as a result of other factors such as
material handling carelessness which might have compromised P. cinnamomi cultures.
Curiously, P. cinnamomi is able to grow at 3 mg/mL plus DMSO 5 % during the ethyl acetate
fraction conditioned treatments. The fact that the oomycete is able to grow at this comparatively
similar concentration with the presence of DMSO seems to support the aforementioned
statement. Previous studies conducted with various solanaceous plants report similar results to
the ones observed in S. betaceum fractions during the present study. Muto et al. (2006) reports
that nightshade (Solanum nigrum) roots n-butanol extract is effective at inhibiting conidial
germination in the fungal pathogen Alternaria brassicicola, and studies carried out by Khan et
al. (2011) conducted with night-blooming jasmine (Cestrum nocturnum) shows that the whole
plant n-butanol extract have an high antimicrobial activity against several fungal and bacterial
pathogenic strains. No P. cinnamomi growth inhibition was noted during the remaining
standard treatments conducted with the S. betaceum ethanol, n-hexane, chloroform and aqueous
fractions, so no further conclusions can be made about the mentioned fractions MIC/MLC. The
lack of activity of the ethanol extract may be explained by the concentration of the extract
compounds, which were too diluted in the ethanol extract and became more concentrated in the
n-butanol fraction. Additionally, the aqueous fraction EFAA results reveals that the fraction is
not very effective at reducing P. cinnamomi HGA, especially when compared to the other S.
betaceum fraction of n-butanol tested in EFAA. The aqueous fraction is approximately half as
effective as the n-butanol fraction regarding P. cinnamomi growth reduction (GR), with a
minimum reduction of 8.60 ± 10.35 % in the 1.25 mg/mL treatment and a maximum reduction
of 53.00 ± 3.41 % in the 10 mg/mL treatment. It is important to highlight that the aqueous
fraction concentrations tested during EFAA present little statistically significant differences
57
amongst themselves and the 1.25 and 5 mg/mL treatments shows highly variable results, so
much so that one of the 1.25 mg/mL treatment replicates actually increases P. cinnamomi HGA
by 6.60 % when compared to the control HGA mean. This outlier result may be due to various
reasons, but the difference in fraction plus growth medium mixture can be excluded, as the
treatment growth solutions were prepared prior to their addition and distribution to the Petri
dishes. A possible explanation may be the hormesis effect. Hormesis is highly specific between
the active agent and the exposed species (Calabrese et al., 2019) and its effects are apparent and
widely reported in oomycetes and fungi alike. For instance, studies by Kato et al. (1990) and
Zhang et al. (1997) report that the common fungicide hymexazol stimulates growth at low
concentrations in Phytophthora spp.. Likewise, studies by Fenn & Coffey (1984) conducted
with phosphorus acid show that the compound boosts the growth of Pythium ultimum and
Pythium myriotylum at low doses. In present study, the exposure to low concentrations of the
supposedly inhibitory aqueous fraction may trigger a compensatory stress response in P.
cinnamomi and promote its growth, with further testing being needed. In contrast with the
previously assessed n-butanol fraction, a higher concentration of polysaccharides than
polyphenols can be observed in the aqueous fraction, with a seemingly similar concentration of
terpenoid compounds. A substantial decrease in ester compounds is also observed, which
suggests a reduction in the fraction lipid content. Both chemical composition changes are to be
expected, given that polysaccharides and terpenoids are relatively soluble in water (due to
hydrogen bonding), while polyphenols are only sparingly soluble and lipids are hydrophobic,
both more soluble in non-polar solvents such as n-butanol. This reduction in polyphenols may
be influencing the inhibitory activity of the aqueous fraction in relation with the n-butanol
fraction, again suggesting that P. cinnamomi is likely showing a higher sensitivity towards the
polyphenol content rather than the polysaccharide compounds.
Results analysis - C. album fractions
The standard treatments conducted using C. album extract fractions held much more
interesting results. We were able to assess most of the tested fractions MIC/MLC and draw
clearer conclusions regarding the fractions inhibitory capacity. Unfortunately, several
conflicting results arose from PTG tests when compared to the corresponding MAA treatments
results, which will be discussed throughout. The n-hexane fraction inhibits P. cinnamomi
growth starting at 1.75 mg/mL, indicating that the fraction MIC may possibly be 1.75 mg/mL.
However, this conclusion cannot be fully supported, as the lethality of this concentration was
not assessed with additional PTG tests. The treatments conducted with the chloroform fraction
58
reveal that 1.75 mg/mL is lethal to P. cinnamomi and likely is the chloroform fraction MLC,
considering that the material treated at 1.75 mg/mL and above failed to grow during PTG tests.
This indicates that the chloroform MIC has to be between 1.50-1.75 mg/mL. Curiously, the
oomycete material treated at 1.50 mg/mL, which did grow during the MAA treatments, failed
to grow in the succeeding PTG tests. This might be a result of carelessness in material handling
or rinsing during the oomycete transfer (e.g.: tongs being too hot; ineffective fraction rinsing)
from the multiwell to the Petri dish in which PTG were conducted. The ethyl acetate fraction
inhibits P. cinnamomi growth starting at 1.25 mg/mL. This inhibited oomycete growth (except
the 1.50 mg/mL treated material) was subsequently subject to PTG tests in which it also failed
to grow, concluding that 1.25 mg/mL is the ethyl acetate fraction MLC and that the fraction
MIC is within 1.00-1.25 mg/mL. Not many conclusions can be drawn from the C. album n-
butanol fraction treatments. Despite its high solubility in water, which allows the testing of
relatively high concentrations, the mixture of fraction plus PDA medium form a dense
precipitate in the solution over time starting at 10.25 mg/mL, making it impossible to observe
P. cinnamomi hyphae growth and draw results from these treatments. Nonetheless, further PTG
tests were conducted with oomycete material treated with the n-butanol fraction between 10-
10.75 mg/mL. This testing brought contradicting results, with oomycete material treated with
1.75 mg/mL successfully growing, but material treated with 1.5 mg/mL failing to do so. Yet
again, this irregular result is likely to be a result of P. cinnamomi material handling sloppiness.
The only conclusion that can be drawn from the n-butanol MAA treatments is that, despite the
lack of observable results, 10.75 mg/mL cannot be the fraction MLC, as P. cinnamomi material
treated at this concentration successfully grew in PTG tests. The n-butanol fraction was also
used in EFAA treatments, where we obtained interesting results. From all the EFAA fractions
tested, it was the most effective fraction at reducing P. cinnamomi HGA, with an impressive
reduction of 65.81 ± 4.41 % in the lowest concentration tested of 1.25 mg/mL treatment and
completely inhibiting P. cinnamomi growth in the 10 mg/mL treatment. These are interesting
results, as the concentration of 10 mg/mL failed to inhibit the pathogen growth during the MAA
treatments but successfully inhibited it during the EFAA treatments. Only the growth medium
used differ in the different assays, with PDB being used in MAA and PDA being used in EFAA.
Being the only difference amongst treatments, the growth medium seems to influence P.
cinnamomi ability to grow and may have synergistic properties with the extract, being further
research needed to confirm this assessment. Additionally, the C. album n-butanol fraction was
tested in conditioned treatments between 11-18 mg/mL without PDB, successfully inhibiting
oomycete growth in all treatments. Further PTG tests conducted with P. cinnamomi material
59
treated at 11 and 12 mg/mL without PDB reveal that these concentrations are lethal to the
oomycete. On a side note, no precipitate is formed during the C. album conditioned n-butanol
fraction treatments without PDB, akin to what occurred in the standard treatments at the same
tested concentrations. This is likely due to the absence of PDB medium, which forms the
precipitate by reacting with the n-butanol fraction. The aqueous fraction did not show any
relevant results, failing to inhibit P. cinnamomi growth even at concentrations as high as 18
mg/mL. This result suggest that the aqueous fraction MIC is far from being reached. The
fraction was also used during EFAA tests, where it showed modest results at 5 and 10 mg/mL,
with an oomycete HGA reduction of 32.94 ± 12.76 % and 57.89 ± 2.58 %, respectively.
Surprisingly, the aqueous fraction boosts P. cinnamomi HGA in all the 1.25 mg/mL replicates
by 9.02 ± 2.37 % increase, and in one 2.5 mg/mL replicate, which increase HGA by 13.00 %.
The possible explanation to this growth stimulation may be the hormesis effect, which was
previously explained in similar results observed in the S. betaceum aqueous fraction (see Results
analysis - S. betaceum fractions). Inhibiting P. cinnamomi growth starting at 0.7 mg/mL, the
ethanol extract has the lowest inhibitory concentration of all tested fractions in the entire present
study, showing impressive results. The oomycete material which failed to grow was subject to
PTG tests, in which it also failed to grow, suggesting that 0.7 mg/mL is the MLC of the ethanol
extract and that its MIC must be between 0.625-0.7 mg/mL. Oomycete material treated at 1.25
and 2.5 mg/mL in the MAA did grow during PTG tests. These two outliers are particularly
strange, as the usual outlier results occur when oomycete material fails to grow in between
successful results and can be reasonably justified with material handling carelessness. Most of
the C. album fractions possess acids (presence of negatively charged carboxyl groups), aromatic
compounds and ester compounds in their composition, which are likely the compounds
responsible for the inhibition of P. cinnamomi growth. The anti-oomycete activity of various
acid compounds is well documented against Phytophthora spp. (e.g.: Lee et al., 2004; Son et
al., 2008), as well as in aromatic compounds (e.g.: Montenegro & Madrid, 2019; McKee et al.,
2020). Due to the non-specific chemical characterization conducted in this study, no further
assessments can be made of which compounds class are present in the C. album fractions. The
detected acid may be related to a plethora of different compounds which possess COO- groups,
and the aromatic compounds can be related to phenolic compounds or alkaloids (aromatic ring),
which are highly different in terms of their bioactivity, with further research being needed to
identify the specific compounds present. The detection of triterpenoids in several of the C.
album fractions also corroborates with the detection of aromatic compounds.
60
The C. album n-butanol and aqueous fractions showed a relatively small anti-oomycete
activity during the MAA treatments, when compared to the remaining tested fractions.
Nonetheless, the n-butanol fraction inhibition shown during the EFAA treatments is
astonishing, effectively halting P. cinnamomi growth at 10 mg/mL. The fraction possesses a
high concentration of aromatic compounds, polysaccharides and triterpenoids, akin to most of
the remaining C. album fractions. Further tests should had been conducted using the other
fractions to fully assess the compounds anti-oomycete activity, but the lack of crude extract and
the fractionation yield conditioned the number of possible tests and fraction material we could
use during the present study.
DMSO and growth medium influence on P. cinnamomi
The conditioned treatments conducted with both species fractions seem to indicate that
the PDB has substantially more influence in oomycete growth than the DMSO, as the PDB
presence allow P. cinnamomi to grow in treatments with up to a 10x greater DMSO percentage.
This is to be expected as the PDB medium supply nutrients which sustain the oomycete growth.
P. cinnamomi is capable of growing in several of the conditioned treatments conducted with
DMSO 5 %, but the control treatments show that DMSO 5 % by itself is lethal to P. cinnamomi.
This observation suggests that DMSO and our extract fractions may have synergistic properties
amongst themselves, but this assessment cannot be fully supported by the present study results
alone, being further research needed.
Taken together, these results show that there are substantial differences between the two
species fractions regarding the inhibition of P. cinnamomi. C. album fractions are significantly
more effective in the control of the pathogen growth, especially the ethanolic initial extract,
which is seemingly as effective at low concentrations as the fungicide Aliette Flash. The C.
album ethanol extract successfully halts P. cinnamomi growth at 0.7 mg/mL, similarly to the
lowest tested concentration of Aliette Flash of 0.75 mg/mL. It is relevant to point out that the
determined MIC/MLC of our extract fraction may not be the exact concentrations, as they may
be found between the known tested concentrations in which P. cinnamomi grew and failed to
do so, such as the C. album n-hexane fraction MIC, that may actually be found within 1.50-
1.75 mg/mL. To precisely find the MIC/MLC of a given fraction, exhaustive concentration
honing tests must be conducted. Furthermore, additional precise treatments can always be made
in accordance with preceding results, resulting in a seemingly endless number of possible tests
61
in search of the exact MIC/MLC concentrations. The limited fraction material available to test
in the present study strictly conditions the number of testing that can be handled, so
concentration honing tests had to be reasonably conducted. The Portuguese-crowberry belongs
to the Ericaceae family, a particularly susceptible family to P. cinnamomi which causes a vast
array of root diseases in the family species (Moreira & Martins, 2005; Robin et al., 2012;
Newhook, 2020), thus the increased extract effectiveness against P. cinnamomi is an interesting
and unexpected result. Recent studies conducted with Arbutus unedo, another Ericaceae, by
Martins et al., (2021) report similar results using the species leaf extracts, proving its
effectiveness in reducing P. cinnamomi growth.
Trichomes
The identification of trichomes in tamarillo was strongly based on trichome
morphological characteristics described by Luckwill (1943), which thoroughly described and
identified the trichome morphology of Lycopersicon. Luckwill (1943) classifies Lycopersicon
trichomes as type I-VII, being types I, IV, VI and VII glandular trichomes and types II, III, V
non-glandular trichomes. The trichome types observed in this study share many morphological
similarities and can be compared with the trichomes types described by Luckwill (1943). Type
A1, A2 and A3 glandular trichomes are similar to the type VI, VII and IV respectively, and the
type B non-glandular trichomes resemble type III trichomes. S. betaceum presents much more
non-glandular trichomes than glandular trichomes. The glandular to non-glandular trichomes
ratio seems to be largely dependent of abiotic and biotic environment factors such as water
availability (Lauter & Munns, 1986). Plants need to spend water in the production and
maintenance of glandular trichomes and their exudates, which may be disadvantageous in hotter
environments (Lauter & Munns, 1986; Van Dam et al., 1999), and favor the production of non-
glandular trichomes. The production of non-glandular trichomes can also be related to
herbivorous insects population density (Gibson, 1979). In all the tamarillo genotypes observed,
the indumentum is significantly denser in the abaxial leaf page and along the leaves midrib and
veins. This density difference between the adaxial and abaxial leaf pages may be due to the
trichomes role in stomata protection against damaging UV radiation (Grammatikopoulos et al.,
1994) or transpiration control. Brewer et al. (1991) and Fernández et al. (2014b) report that
higher trichome density in the abaxial leaf surface result in a lower surface water retention,
which decreases stomatal occlusion and promotes CO2 leaf exchanges in Glycine max and
Quercus ilex leaves, respectively. Further studies should be conducted to access the role of S.
betaceum trichomes in plant defense and water retention. C9 and C12 varieties have much less
62
trichomes than the red and orange-type varieties. This observation was expected, as the leaves
from the in vitro genotypes are much younger than the leaves observed from the red and orange-
type, and the in vitro plantlets are not as exposed to biotic and abiotic factors, which stimulate
trichome production. The A3 type found in the in vitro genotypes may be a result of mutations
that condition trichome development or deviations in the formation of type A1 trichomes. We
cannot assume with certainty that this type of trichomes have a single secretory cell, as a bundle
of cells may be encapsuled in a membrane, visually appearing as a single cell. In all four S.
betaceum variants trichomes were significantly denser in the abaxial leaf surface and along the
midrib and nervures. Three types of trichomes were observed in all tamarillo varieties, one type
of needle-like (or apiculate) multicellular non-glandular trichomes, and two types of
multicellular glandular trichomes, one being long and the other being short, both with four
coplanar conspicuous secretory cells. These types of trichomes can be observed in different
stages of maturation, leading to morphological differences such as size and number of cells.
These observations go in accordance with observations made by Mahlberg (1985).
C. album presents a large number of long non-glandular trichomes and only a small
number of glandular trichomes are observed. The presence of a thick mucilage at the abaxial
surface suggests that a substantial amount of glandular trichomes should be present, from which
the mucilage would secrete from, but only a very small number of glandular trichomes were
observed, possibly due to the thick mucilage covering them. The adaxial surface is completely
glabrous, where no trichomes are present. From a personal observation, the C1 type trichomes
are morphologically similar to the D type trichomes. The D type trichomes are long non-
glandular trichomes, seemingly unicellular. The C1 type are long glandular multicellular
trichomes and the C2 type are short glandular multicellular trichomes. These observations go
in accordance with observations made in Antunes et al. (2018). The compounds tested in the
present study may be present either in the leaf cells themselves or in the thick mucilage present,
with further studies being needed to fully assess the compounds origin.
63
5. CONCLUSIONS AND FUTURE PERSPECTIVES
The study of the activity of plant secondary metabolites such as plant stress tolerance give
way to the discovery of potential new applications to these compounds and generate an
increasing interest in these compounds research and production. The promising results obtained
provide a first approach and contribute to future research concerning the identification and
purification of the tested fractions compounds which can be further applied in selective
breeding programs to develop more resistant cultivars.. The present study shows that tamarillo
and crowberry leaves may be useful materials to produce natural plant products to control P.
cinnamomi infections and its associated diseases, and possibly contribute to the discovery of an
environmentally-friendly alternative to phosphites and their derivatives pesticides, which
currently are the best (and worst) treatment solution to P. cinnamomi control.
Given the present study results obtained with the C. album crude ethanol extract and the
n-butanol fraction, it would be of interest to further research these specific fractions
composition and assess P. cinnamomi sensitivity towards these compounds. Moreover, the
studied fractions effectiveness should also be subsequentially tested in vivo, which also let us
evaluate if the fractions are toxic to the plant organisms subjected to the treatment and if
external factors not present in in vitro testing influence the fractions effectiveness.
64
65
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79
APPENDIXES
Appendix 1. Multiwell activity assay #1 growth results. Green – Growth; Red – No growth.
Appendix 2. Multiwell activity assay #2 growth results. ** A dense precipitate was found and hyphal
growth could not be observed. Green – Growth; Red – No growth.
Appendix 3. Multiwell activity assay #3 growth results. Green – Growth; Red – No growth.
Appendix 4. Multiwell activity assay #4 growth results. Green – Growth; Red – No growth.
Appendix 5. Multiwell activity assay #5 growth results. * A dense precipitate was found and hyphal
growth could not be observed. Green – Growth; Red – No growth.
Appendix 6. Multiwell activity assay #6 growth results. * Control replicate did not grow.
Green – Growth; Red – No growth.
Treatment
C (mg/mL)
Area (cm2)
Tamarillo
n-Butanol fraction
(4-BUT)
10
5,381
5,499
6,003
5
6,582
8,133
8,792
2.5
10,389
14,851
13,329
1.25
21,063
21,336
19,574
Tamarillo
Aqueous fraction
(5-AQU)
10
11,113
9,304
10,723
5
14,479
17,561
13,876
2.5
18,897
17,713
17,595
1.25
24,318
20,852
18,576
Crowberry
n-Butanol fraction
(4-BUT)
10
0
0
0
5
2,424
2,000
2,624
2.5
3,998
5,157
4,279
1.25
6,888
9,331
7,830
Crowberry
Aqueous fraction
(5-AQU)
10
9,608
9,913
8,539
5
15,327
13,978
15,299
2.5
25,777
17,663
22,474
1.25
25,971
24,871
24,786
PDA
-
22,679
22,639
23,121
PDA 2x
-
16,054
18,789
Appendix 7. Extract fraction activity assay (EFAA) growth area results.
13,750
C (mg/mL)
TG (cm2)
GR (cm2)
GR (%)
GRmean (%)
StdDev
1.25
21.063
1.750
7.671
7.671
3.39
21.336
1.477
6.474
19.574
3.239
14.198
2.5
10.389
12.424
54.460
41.573
8.12
14.851
7.962
34.901
13.329
9.484
41.573
5
6.582
16.231
71.148
64.349
4.06
8.133
14.680
64.349
8.792
14.021
61.461
10
5.381
17.432
76.413
75.895
1.18
5.499
17.314
75.895
6.003
16.810
73.686
SS
df
MS
F
P-value
Fcrit
Between groups
7688.3297
3
2562.7766
71.6997
0.00000401
4.0662
Within groups
285.9454
8
35.7432
-
-
-
Total
7974.2751
11
-
-
-
-
Groups
1.25 vs 2.5
1.25 vs 5
1.25 vs 10
2.5 vs 5
2.5 vs 10
5 vs 10
1.25
2.5
1.25
5
1.25
10
2.5
5
2.5
10
5
10
Average
9.4478
43.6447
9.4478
65.6526
9.4478
75.3313
43.6447
65.6526
43.6447
75.3313
65.6526
75.3313
Variance
17.2814
98.8585
17.2814
24.7358
17.2814
2.0970
98.8585
24.7358
98.8585
2.0970
24.7358
2.0970
N
3
3
3
3
3
3
3
3
3
3
3
3
P. variance
58.0700
-
21.0086
-
9.6892
-
61.7971
-
50.4778
-
13.4164
-
H. mean
0
-
0
-
0
-
0
-
0
-
0
-
df
4
-
4
-
4
-
4
-
4
-
4
-
t Stat
-5.4961
-
-15.0183
-
-25.9225
-
-3.4288
-
-5.4622
-
-3.2363
-
P one-tail
0.0027
-
0.0001
-
0.0000
-
0.0133
-
0.0027
-
0.0159
-
tcrit one-tail
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
P two-tail
0.0053
-
0.0001
-
0.0000
-
0.0266
-
0.0055
-
0.0318
-
tcrit two-tail
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
Bonferroni correction: α = p significance/nº comparisons = 0.05/6 = 0.00833
Appendix 9. S. betaceum 4-BUT EFAA (one-way ANOVA; p=0.05).
Appendix 10. S. betaceum 4-BUT EFAA post hoc Tuckey test (p=0.05). Significative or non-
significative differences between groups are marked as green and red, respectively.
Appendix 8. S. betaceum 4-BUT EFAA treatment growth (TG) and growth reduction (GR).
C (mg/mL)
TG (cm2)
GR (cm2)
GR (%)
GRmean (%)
StdDev
1.25
24.318
-1.505
-6.597
8.596
10.35
20.852
1.961
8.596
18.576
4.237
18.573
2.5
18.897
3.916
17.166
22.356
2.58
17.713
5.100
22.356
17.595
5.218
22.873
5
14.479
8.334
36.532
36.532
7.07
17.561
5.252
23.022
13.876
8.937
39.175
10
11.113
11.700
51.287
52.996
3.41
9.304
13.509
59.216
10.723
12.090
52.996
SS
df
MS
F
P-value
Fcrit
Between groups
3668.6164
3
1222.8721
18.59197
0.00057789
4.0662
Within groups
526.1938
8
65.7742
-
-
-
Total
4194.8102
11
-
-
-
-
Groups
1.25 vs 2.5
1.25 vs 5
1.25 vs 10
2.5 vs 5
2.5 vs 10
5 vs 10
1.25
2.5
1.25
5
1.25
10
2.5
5
2.5
10
5
10
Average
6.8572
20.7981
6.8572
32.9096
6.8572
54.4996
20.7981
32.9096
20.7981
54.4996
32.9096
54.4996
Variance
160.6480
9.9628
160.6480
75.0707
160.6480
17.4154
9.9628
75.0707
9.9628
17.4154
75.0707
17.4154
N
3
3
3
3
3
3
3
3
3
3
3
3
P. variance
85.3054
-
117.8593
-
89.0317
-
42.5167
-
13.6891
-
46.2431
-
H. mean
0
-
0
-
0
-
0
-
0
-
0
-
df
4
-
4
-
4
-
4
-
4
-
4
-
t Stat
-1.8486
-
-2.9391
-
-6.1840
-
-2.2749
-
-11.1560
-
-3.8884
-
P one-tail
0.0691
-
0.0212
-
0.0017
-
0.0426
-
0.0002
-
0.0089
-
tcrit one-tail
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
P two-tail
0.1382
-
0.0424
-
0.0035
-
0.0853
-
0.0004
-
0.0177
-
tcrit two-tail
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
Bonferroni correction: α = p significance/nº comparisons = 0.05/6 = 0.00833
Appendix 12. S. betaceum 5-AQU EFAA (one-way ANOVA; p=0.05).
Appendix 13. S. betaceum 5-AQU EFAA post hoc Tuckey test (p=0.05). Significative or non-
significative differences between groups are marked as green and red, respectively.
Appendix 11. S. betaceum 5-AQU EFAA treatment growth (TG) and growth reduction (GR).
C (mg/mL)
TG (cm2)
GR (cm2)
GR (%)
GRmean (%)
StdDev
1.25
6.888
15.925
69.807
65.677
4.41
9.331
13.482
59.098
7.83
14.983
65.677
2.5
3.998
18.815
82.475
81.243
2.16
5.157
17.656
77.394
4.279
18.534
81.243
5
2.424
20.389
89.374
89.374
1.14
2
20.813
91.233
2.624
20.189
88.498
10
0
22.813
100
100
0
0
22.813
100
0
22.813
100
SS
df
MS
F
P-value
Fcrit
Between groups
2003.1307
3
667.7102
70.01962
0.00000439
4.0662
Within groups
76.2884
8
9.5360
-
-
-
Total
2079.4190
11
-
-
-
-
Groups
1.25 vs 2.5
1.25 vs 5
1.25 vs 10
2.5 vs 5
2.5 vs 10
5 vs 10
1.25
2.5
1.25
5
1.25
10
2.5
5
2.5
10
5
10
Average
64.8607
80.3708
64.8607
89.7018
64.8607
100
80.3708
89.7018
80.3708
100
89.7018
100
Variance
29.1700
7.0234
29.1700
1.9508
29.1700
0
7.0234
1.9508
7.0234
0
1.9508
0
N
3
3
3
3
3
3
3
3
3
3
3
3
P. variance
18.0967
-
15.5604
-
14.5850
-
4.4871
-
3.5117
-
0.9754
-
H. mean
0
-
0
-
0
-
0
-
0
-
0
-
df
4
-
4
-
4
-
4
-
4
-
4
-
t Stat
-4.4654
-
-7.7127
-
-11.2690
-
-5.3950
-
-12.8289
-
-12.7708
-
P one-tail
0.0056
-
0.0008
-
0.0002
-
0.0029
-
0.0001
-
0.0001
-
tcrit one-tail
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
P two-tail
0.0111
-
0.0015
-
0.0004
-
0.0057
-
0.0002
-
0.0002
-
tcrit two-tail
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
Bonferroni correction: α = p significance/nº comparisons = 0.05/6 = 0.00833
Appendix 15. C. album 4-BUT EFAA (one-way ANOVA; p=0.05).
Appendix 16. C. album 4-BUT EFAA post hoc Tuckey test (p=0.05). Significative or non-
significative differences between groups are marked as green and red, respectively.
Appendix 14. C. album 4-BUT EFAA treatment growth (TG) and growth reduction (GR).
C (mg/mL)
TG (cm2)
GR (cm2)
GR (%)
GRmean (%)
StdDev
1.25
25.971
-3.158
-13.843
-9.021
2.37
24.871
-2.058
-9.021
24.786
-1.973
-8.649
2.5
25.777
-2.964
-12.993
1.486
14.60
17.663
5.150
22.575
22.474
0.339
1.486
5
15.327
7.486
32.815
32.937
2.76
13.978
8.835
38.728
15.299
7.514
32.937
10
9.608
13.205
57.884
57.884
2.58
9.913
12.900
56.547
8.539
14.274
62.570
SS
df
MS
F
P-value
Fcrit
Between groups
8775.2421
3
2925.0807
33.45633
0.00007089
4.0662
Within groups
699.4385
8
87.4298
-
-
-
Total
9474.6806
11
-
-
-
-
Groups
1.25 vs 2.5
1.25 vs 5
1.25 vs 10
2.5 vs 5
2.5 vs 10
5 vs 10
1.25
2.5
1.25
5
1.25
10
2.5
5
2.5
10
5
10
Average
-10.5042
3.6894
-10.5042
34.8266
-10.5042
59.0000
3.6894
34.8266
3.6894
59.0000
34.8266
59.0000
Variance
8.3951
319.9020
8.3951
11.4188
8.3951
10.0034
319.9020
11.4188
319.9020
10.0034
11.4188
10.0034
N
3
3
3
3
3
3
3
3
3
3
3
3
P. variance
164.1485
-
9.9069
-
9.1993
-
165.6604
-
164.9527
-
10.7111
-
H. mean
0
-
0
-
0
-
0
-
0
-
0
-
df
4
-
4
-
4
-
4
-
4
-
4
-
t Stat
-1.3568
-
-17.6388
-
-28.0660
-
-2.9629
-
-5.2744
-
-9.0462
-
P one-tail
0.1232
-
0.0000
-
0.0000
-
0.0207
-
0.0031
-
0.0004
-
tcrit one-tail
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
2.1318
-
P two-tail
0.2464
-
0.0001
-
0.0000
-
0.0414
-
0.0062
-
0.0008
-
tcrit two-tail
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
2.7764
-
Bonferroni correction: α = p significance/nº comparisons = 0.05/6 = 0.00833
Appendix 18. C. album 5-AQU EFAA (one-way ANOVA; p=0.05).
Appendix 19. C. album 5-AQU EFAA post hoc Tuckey test (p=0.05). Significative or non-
significative differences between groups are marked as green and red, respectively.
Appendix 17. C. album 5-AQU EFAA treatment growth (TG) and growth reduction (GR).
Appendix 20. FTIR absorbance spectrum of S. betaceum ethanol extract.
Appendix 21. FTIR absorbance spectrum of S. betaceum n-hexane fraction.
Appendix 22. FTIR absorbance spectrum of S. betaceum chloroform fraction.
Appendix 23. FTIR absorbance spectrum of S. betaceum ethyl acetate fraction.
Appendix 24. FTIR absorbance spectrum of S. betaceum biomass.
Appendix 25. FTIR absorbance spectrum of C. album biomass.
