Glomeromycotean associations in liverworts: A molecular cellular and taxonomic analysis

Abstract

Liverworts form endophytic associations with fungi that mirror mycorrhizal associations in tracheophytes. Here we report a worldwide survey of liverwort associations with glomeromycotean fungi (GAs), together with a comparative molecular and cellular analysis in representative species. Liverwort GAs are circumscribed by a basal assemblage embracing the Haplomitriopsida, the Marchantiopsida (except a few mostly derived clades), and part of the Metzgeriidae. Fungal endophytes from Haplomitrium, Conocephalum, Fossombronia, and Pellia were related to Glomus Group A, while the endophyte from Monoclea was related to Acaulospora. An isolate of G. mosseae colonized axenic thalli of Conocephalum, producing an association similar to that in the wild. Fungal colonization in marchantialean liverworts suppressed cell wall autofluorescence and elicited the deposition of a new wall layer that specifically bound the monoclonal antibody CCRC-M1 against fucosylated side groups associated with xyloglucan and rhamnogalacturonan I. The interfacial material covering the intracellular fungus contained the same epitopes present in host cell walls. The taxonomic distribution and cytology of liverwort GAs suggest an ancient origin and multiple more recent losses, but the occurence in widely separated liverwort taxa of fungi related to glomeromycotean lineages that form arbuscular mycorrhizas in tracheophytes, notably the Glomus Group A, is better explained by host shifting from tracheophytes to liverworts.

Full text

GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS:
A MOLECULAR,CELLULAR,AND TAXONOMIC ANALYSIS
1
ROBERTO LIGRONE,
2,5
ANNA CARAFA,
2
ERICA LUMINI,
3
VALERIA BIANCIOTTO,
3
PAOLA BONFANTE,
3
AND JEFFREY G. DUCKETT
4
2
Dipartimento di Scienze ambientali, Seconda Universita` di Napoli, via A. Vivaldi 43, I-81100 Caserta, Italy;
3
Dipartimento di Biologia vegetale, Universita` degli Studi di Torino, and Consiglio Nazionale delle Ricerche (CNR),
Istituto per la Protezione delle Piante, Sezione di Torino, Viale P. A. Mattioli 25, I-10125, Torino, Italy; and
4
School of Biological and Chemical Sciences, Queen Mary University of London, Mile End Road, London E1 4NS, UK
Liverworts form endophytic associations with fungi that mirror mycorrhizal associations in tracheophytes. Here we report a
worldwide survey of liverwort associations with glomeromycotean fungi (GAs), together with a comparative molecular and
cellular analysis in representative species. Liverwort GAs are circumscribed by a basal assemblage embracing the
Haplomitriopsida, the Marchantiopsida (except a few mostly derived clades), and part of the Metzgeriidae. Fungal endophytes
from Haplomitrium,Conocephalum,Fossombronia, and Pellia were related to Glomus Group A, while the endophyte from
Monoclea was related to Acaulospora. An isolate of G. mosseae colonized axenic thalli of Conocephalum, producing an
association similar to that in the wild. Fungal colonization in marchantialean liverworts suppressed cell wall autofluorescence and
elicited the deposition of a new wall layer that specifically bound the monoclonal antibody CCRC-M1 against fucosylated side
groups associated with xyloglucan and rhamnogalacturonan I. The interfacial material covering the intracellular fungus contained
the same epitopes present in host cell walls. The taxonomic distribution and cytology of liverwort GAs suggest an ancient origin
and multiple more recent losses, but the occurence in widely separated liverwort taxa of fungi related to glomeromycotean
lineages that form arbuscular mycorrhizas in tracheophytes, notably the Glomus Group A, is better explained by host shifting from
tracheophytes to liverworts.
Key words: arbuscular mycorrhizas; cell walls; DNA sequencing; Glomeromycota; immunocytochemistry; liverworts;
symbiosis; ultrastructure.
The establishment of biotrophic associations with fungi is
considered a major factor involved in the colonization of
terrestrial habitats by phototrophic organisms (Selosse and Le
Tacon, 1998). It is assumed that the common ancestor to the
Glomeromycota, Ascomycota, and Basidiomycota originated
after the appearance of land plants (Berbee and Taylor, 2007)
and that the association with glomeromycotean fungi, to form
the so-called arbuscular mycorrhizas (AMs), is a plesiomorphy
(primitive character) in the tracheophytes. Already present in
Siluro-Devonian fossils of protracheophytes and still occurring
in the majority of present-day tracheophytes (Selosse and Le
Tacon, 1998; Wang and Qiu, 2006), the AMs have been
replaced by associations with basidio- or ascomycetes in
several derived lineages of higher plants (Wang and Qiu, 2006;
Berbee and Taylor, 2007). Endophytic fungal associations not
only occur in tracheophytes but also in the gametophytes of
liverworts and hornworts, while they appear to be absent in
mosses (Read et al., 2000; Renzaglia et al., 2007).
The fungal associations in members of the Marchantiopsida
(complex thalloid liverworts) and Metzgeriidae (simple thalloid
liverworts) are cytologically similar to AMs (Strullu et al.,
1981; Pocock and Duckett, 1984; Ligrone and Lopes, 1989;
Ligrone and Duckett, 1994). Similar associations have also
been described in Haplomitrium and Treubia (Carafa et al.,
2003; Duckett et al., 2006a), two taxa recently placed in a clade
that is sister to all other liverworts (Forrest and Crandall-
Stotler, 2004, 2005; Heinrichs et al., 2005; Forrest et al., 2006).
With the application of molecular techniques, the fungal
symbiont in Marchantia foliacea has been identified as
belonging to the glomeromycotean genus Glomus, group A
(Russell and Bulman, 2005). An assemblage of simple thalloid
liverworts and the leafy liverworts (Jungermanniidae) form a
diversity of endophytic associations with asco- or basidiomy-
cetes or are fungus-free (Kottke et al., 2003; Nebel et al., 2004;
Duckett et al., 2006b).
With reference to the topology of liverwort phylogeny as
revealed by recent molecular work (Davis, 2004; Forrest and
Crandall-Stotler, 2004, 2005; Heinrichs et al., 2005), it has
been suggested that the association with glomeromycotean
fungi is a plesiomorphy in the liverworts (Nebel et al., 2004;
Kottke and Nebel, 2005). Moreover, considering that the
liverworts are almost unanimously recognized as the earliest-
1
Manuscript received 24 February 2007; revision accepted 16 August
2007.
This work was funded by grants from the Seconda Universita` di Napoli
and Regione Campania, Italy (LR 5, 2003). The research in Torino was
funded by the Biodiversity Project of CNR, Italy. The authors thank M.
Hahn (Complex Carbohydrate Research Center, University of Georgia,
USA) and J. P. Knox (Centre for Plant Sciences, University of Leeds, UK)
for the generous gift of the antibodies used in this study, and V.
Gianinazzi-Pearson (INRA, Dijon, France) for supplying the spores of G.
mosseae and G. clarum. The authors also thank K. Renzaglia, the staff at
the IMAGE Center (Southern Illinois University), and the staff at the
CISME (University of Naples ‘‘Federico I,’’ Italy) for laboratory and
electron microscopy facilities; K. Pell (QMUL) for technical assistance;
the Department of Plant and Microbial Sciences, University of Canterbury,
Christchurch, New Zealand, for laboratory facilities; the New Zealand
Department of Conservation for granting collecting permits; and B.
Butterfield (University of Canterbury) and D. Glennie (Landcare, Lincoln,
New Zealand) for their help in the collection of the specimens used in this
study. J.G.D. was supported by an overseas travel grant from the Royal
Society (UK) in New Zealand and by a DEFRA Darwin Initiative grant in
Chile. R.L. was supported by a grant from CNR (Italy) in New Zealand.
5
Author for correspondence (e-mail: roberto.ligrone@unina2.it)
American Journal of Botany 94(11): 1756–1777. 2007.

divergent clade in the phyletic tree of land plants (Nickrent et
al., 2000; Dombrovska and Qiu, 2004; Groth-Malonek et al.,
2005; Qiu et al., 2006) and that the Glomeromycota are basal to
the other mycorrhiza-forming fungi (Schu¨ßler et al., 2001;
James et al., 2006), it has been suggested that glomeromyco-
tean associations (GAs) in liverworts predated the arbuscular
mycorrhizas in vascular plants (Nebel et al., 2004; Kottke and
Nebel, 2005; Duckett et al., 2006a; Wang and Qiu, 2006). An
alternative scenario, i.e., secondary host shift of glomeromy-
cotean symbionts from tracheophytes to liverworts, has been
considered by Selosse (2005), mainly on the basis of Russell
and Bulman’s (2005) identification of the fungal endophyte of
Marchantia paleacea as a member of the Glomus Group A,
i.e., a derived group in the phyletic tree of Glomeromycota
(Schu¨ßler et al., 2001).
In spite of the growing interest in fungus–liverwort
associations in recent years, current information on their
cytology and physiology is remarkably sparse. In particular, as
concerns putative GAs, the information available for most of
the taxa reported by Nebel et al. (2004) is from light
microscopy and generally does not go beyond the notion of
the presence/absence of fungal endophytes tentatively referred
to as glomero, basidio- or ascomycetes. Owing to the small
number of taxa investigated in detail to date, it is impossible to
reach any general conclusions about the cytology of putative
GAs in liverworts.
The general aim of this long-standing investigation was to
provide an exhaustive survey of the biology of GAs in
liverworts and specifically to (1) identify the fungal endophytes
through molecular analysis in selected liverwort taxa; (2)
determine the taxonomic and geographical distribution of GAs
in liverworts through a morphological (light and electron
microscopy) analysis of taxa collected worldwide; (3) inves-
tigate the level of cellular compatibility between liverworts and
fungi through a detailed immunocytochemical analysis of their
contact surfaces; and (4) confirm Koch’s postulates through in
vitro synthesis of GAs from axenic liverwort cultures and
spores of known glomeromycotean fungi. The data presented
are discussed in the context of the origins of GAs in liverworts
and their evolutionary relationships with AMs.
MATERIALS AND METHODS
The liverwort species examined, their taxonomic position, fungal status, and
geographical origin are listed in Table 1. Liverwort taxonomy follows Crandall-
Stotler and Stotler (2000) and Heinrichs et al. (2005). With the exception of few
exceedingly rare species, the diagnosis for fungal status was based on the study
of samples from at least two separate collection sites and from freshly-collected
specimens. At least 20 plants were examined for each sample. For voucher
information of the taxa examined in this study, see the Appendix.
Molecular analysis—Molecular analysis of fungal endophytes was carried
out for the following liverwort species: Haplomitrium chilensis,Conocephalum
conicum, Monoclea gottschei, Fossombronia echinata,Pellia endiviifolia.
Healthy thalli or, in the case of H. chilensis, subterranean mycotrophic axes
(Carafa et al., 2003) were carefully rinsed with distilled water, and colonized
parts were isolated with a razor blade under a dissecting microscope. The
samples, each about 50–100 mg, were surface-sterilized with cloramine T (3%)
and streptomycin (0.3%) followed by two rounds of sonication. A mininum of
two samples for each liverwort species were processed separately.
DNA was extracted using the Dneasy Plant Mini kit (Qiagen, Valencia,
California, USA) according to manufacturer protocols. Partial small-ribosomal-
subunit (SSU) DNA fragments (550 bp) were amplified using the universal
eukaryotic primer NS31 (Simon et al., 1993) and the Glomeromycota-specific
primer AM1 (Helgason et al., 1998). DNA extracts from Glomus mosseae
(BEG12) and Gigaspora rosea (BEG9) isolates were used as positive controls,
while DNA extracts from fungus-free apical parts of the thalli were used as
negative controls.
The PCR reaction was performed in a total volume of 25 lL containing 2 lL
of template solution, 0.2 mM of each dNTP, 10 pmols of each primer, 1 U of
REDTaq DNA polymerase (Sigma, St. Louis, Missouri, USA) and 13REDTaq
Reaction buffer (SIGMA). Amplification was performed in a GeneAmp PCR
system 9700 (PerkinElmer, Waltham, Massachussets, USA) programmed as
follows: 1 33 min at 958C; 35 31 min at 958C, 1 min at 588C, 2 min at 758C;
137 min at 728C. Electrophoretical analysis of the PCR products revealed a
single band of 550 bp. This fragment was purified from gel using the QIAquick
purification kit (QIAGEN), cloned into a pGEM-T Easy Vector (Promega,
Madison, Wisconsin, USA), and then transformed into Escherichia coli JM109
High Efficiency Competent Cells (Promega).
Thirty putatively positive transformant clones (white colonies) from each
liverwort sample were selected manually, and the DNA extracted from each
clone was amplified using the PCR mix and program detailed previously. For
RFLP (restriction fragment length polymorphism) analysis, aliquots of 4 lLof
each PCR amplicon were mixed with 16 lL of digestion mix containing 2.0 lL
buffer 103, 0.2 lL bovine serum albumin, 13.3 lLH
2
O, and 0.5 lL of the
restriction enzyme Hinf IorHsp92II (Promega) for 3 h at 378C. Fragment
patterns were analyzed on agarose gel containing 0.84 % agarose (Sigma) and
1.5% high-resolution agarose (Sigma). One to four PCR amplicons were
sequenced for each restriction pattern and species, using the vector-specific
primers T7 and SP6, at the DNA Sequences Naples Facilities. The sequences
have been deposited in GenBank under the accession numbers reported in
Table 3.
Forward and reverse sequences were analyzed using the program BioLign
4.0.6 (http://en.bio-soft.net/dna/BioLign.html). DNA sequences were compared
to GenBank database using the BLAST algorithm (Altschul et al., 1997) for
identification. Data bank sequences with high homology to our sequences were
included in the data set, using the profile alignment function CLUSTAL W
(Thompson et al., 1994) for multiple alignment. The nearest relatives of each
sequence were inferred with the neighbor-joining algorithm (Saitou and Nei,
1987) and the Kimura two-parameter model (Kimura, 1980), using the PHYLIP
package (Felsenstein, 1989). The confidence of branching was assessed using
1000-bootstrap resampling (Felsenstein, 1985).
Light and electron microscopy—Both fresh and fixed samples were
examined by light microscopy. The visibility of fungal hyphae in rhizoids and
hand-cut sections of the thallus was improved by staining with 0.05% trypan
blue in lactophenol (Ligrone and Lopes, 1989) or 0.05% aniline blue in lactic
acid. Autofluorescence of liverwort cell walls was observed on fresh hand-cut
sections using an excitation filter at 365 nm and a barrier filter with a
transmission cutoff at 397 nm.
Colonized areas of the thallus of fungus-containing specimens were cut into
small pieces under a dissecting microscope and fixed with a mixture of 3%
glutaraldehyde, 1% freshly prepared formaldehyde, and 0.75% tannic acid in
0.04 M piperazine-N,N0–bis(2-ethanesulfonic acid) (PIPES) buffer, pH 7.0, for
2 h at room temperature under gentle vacuum. The samples were then rinsed in
0.08 M PIPES buffer and twice in 0.08 M Na-cacodylate buffer, and postfixed
in 1% OsO
4
in 0.08 M Na-cacodylate buffer, pH 6.7, overnight at 48C.
Following dehydration in a step gradient of ethanol and one step in propylene
oxide at 48C, the samples were slowly infiltrated with Spurr’s resin
(Polysciences, Warrington, Pennsylvania, USA) at 48C, transferred to
polypropylene dishes, and cured at 688C for 24 h. For light microscopy, 0.5-
lm-thick sections of resin-embedded samples were cut with a diamond
histoknife, stained with 0.5% toluidine blue O in 1% Na-tetraborate, and
photographed with a Zeiss Axioskop (Zeiss, Jena, Germany) light microscope
equipped with a Sensicam QE (Applied Scientific Instrumentations, Eugene,
Oregon, USA) digital photocamera. For transmission electron microscopy
(TEM), ultrathin sections were cut with a diamond knife, collected on 300-
mesh uncoated nickel grids, stained with 3% uranyl acetate in 50% methanol
for 15 min and in Reynold’s lead citrate for 10 min, and observed with a Jeol
1200 EX2 (Jeol, Tokyo, Japan) electron microscope.
For scanning electron microscopy (SEM), the samples were cut with a razor
blade and taken through a 1 : 1 ethanol:acetone series to remove the cytoplasm,
osmicated for 48 h in aqueous 2% OsO
4
, and stored in 70% ethanol. The
samples were then dehydrated in anhydrous ethanol and critical point dried
using CO
2
as the transfusion fluid, mounted on stubs, and sputter-coated with
390 nm palladium-gold. The samples were viewed using a Hitachi (Hitachi,
Tokyo, Japan) S570 scanning electron microscope.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

TABLE 1. Fungal associations in liverworts.
Liverwort taxa Fungal status Geographic origin
Haplomitriopsida:
Haplomitriales
Haplomitrium blumei (Nees) R.M. Schust.
a
G Malaysia (2)
H. gibbsiae (Steph.) R.M. Schust.
a
G New Zealand (7), Uganda (1)
H. hookeri (Smith) Nees
a
G UK (5)
H. intermedium Berrie
ac
G Australia (1)
H. ovalifolium R.M. Schust
a
G New Zealand (2)
H. chilensis R.M. Schust.
ac
G Chile (3)
Treubiales
Treubia lacunosa (Colenso) Prosk.
a
G New Zealand (1)
T. lacunosoides Pfeiffer, Frey & Stecha
ac
G New Zealand (6)
T. pygmaea R.M. Schust.
ac
G New Zealand (6)
Marchantiopsida (complex thalloid liverworts):
Blasiales
Blasia pusilla L.
a
— UK (4), USA (1)
Sphaerocarpales
Sphaerocarpos michelii Bellardi
c
— Italy (1), UK (1)
S. texanus Austin
c
— UK (1)
Geothallus tuberosus Campb.
bc
— USA (1)
Riella americana M.Howe & Underwood
c
— USA (1)
Riella helicophylla (Boryet Mont.) Mont.
c
— Greece (1)
Monocleales
Monoclea forsteri Hook.
a
G New Zealand (7)
M. gottschei Lindb.
a
G Chile (2), Mexico (1), Venezuela (1)
Marchantiales
Aytoniaceae
Asterella bachmanii (Steph.) S.W.Arnell
ac
G South Africa (1)
A. muscicola (Steph.) S.W.Arnell
c
G Lesotho (1)
A. wilmsii (Steph.) S.W.Arnell
ac
G South Africa (1)
A. tenera (Mitt.) R.M. Schust.
ac
G New Zealand (4)
A. australis (Hook.f. & Taylor) Verd.
ac
G New Zealand (4)
Cryptomitrium oreoides Perold
c
— Lesotho (2)
Mannia angrogyna (L.) A. Evans
a
— Italy (1)
M. fragrans (Balb.) Frye & L. Clark — China (1), Germany (1)
Plagiochasma exigua (Schiffn.) Steph.
c
G South Africa (2), Lesotho (2)
P. rupestre (J.R.Forst. & G.Forst.) Steph.
a
G (1) South Africa (3), Lesotho (2)
Reboulia hemispherica (L.) Raddi
a
G Italy (2), UK (3), Chile (2)
Wiesnerellaceae
Wiesnerella denudata Schiffn.
b
— (2) Japan (1), Java (1), Nepal (1), Sikkim (1)
Conocephalaceae
Conocephalum conicum (L.) Dumort.
a
G France (1), Italy (2), UK (6), USA (2)
C. salebrosum Szweykowski, Buczkowska & Odrzykoski
c
G UK (2), USA (3)
Lunulariaceae
Lunularia cruciata (L.) Dumort. ex Lindb.
a
G France (1), Italy (2), UK (4)
Marchantiaceae
Bucegia romanica Raddi
bc
— Rumania (2)
Dumortiera hirsuta (Sw.) Nees
a
G Chile (1), France (1), Venezuela (1), UK (1)
Marchantia berteroana Lehm. & Lindb.
c
G Chile (1), Venezuela (1)
M. foliacea Mitt.
c
G Chile (1), New Zealand (2)
M. pappeana Lehm.
a
G Lesotho (2)
M. polymorpha subsp. polymorpha Gottsche, Lindb. & Nees.— UK (4)
M. polymorpha subsp. ruderalis Bischl. & Boisselier — (1) UK (3)
M. polymorpha subsp. montivagans Bischl. & Boisselier
a
G UK (2)
Neohodgsonia mirabilis (H. Perss.) H. Perss.
a
G (3) New Zealand (2)
Preissia quadrata (Scop.) Nees
a
G Italy (1), UK (4)
Monosoleniaceae
Monosolenium tenerum Griffith
c
— Germany (from aquarium) (1), Japan (1)
Peltolepis grandis Lindb.
b
— Norway (1), Russia (Siberia) (1), Switzerland (1)
Cleveaceae
Athalamia hyalina (Sommerf.) S. Hatt.
c
G Italy (1), USA (1)
A. pinguis W. Falc.
bc
G India (1)
Sauteria alpina (Nees) Nees
b
— Switzerland (2)
Exormothecaceae
Aitchisoniella himalayensis Kash.
bc
— India (1)
Exormotheca holstii Steph. — Lesotho (1)
E. pustulosa Mitt.
c
— Lesotho (1)
Stephensoniella brevipedunculata Kash.
bc
— India (1)
Cyathodiaceae —
Cyathodium cavernarum Kunze
c
— Uganda (1)
AMERICAN JOURNAL OF BOTANY [Vol. 94

TABLE 1. Continued.
Liverwort taxa Fungal status Geographic origin
C. foetidissimum Schiffn.
a
— (2) Italy (1)
Corsiniaceae
Corsinia coriandra (Spreng.) Lindb.
a
G (1) Italy (2)
Cronisia fimbriata (Nees) Whittem. & Bischl.
bc
— Brazil (1)
Monocarpaceae
Monocarpus sphaerocarpus Carr
c
— Australia (1)
Targionaceae
Targionia hypophylla L.
a
G France (2), Italy (2), New Zealand (3), UK (2)
Oxymitraceae
Oxymitra incrassata (Broth.) Sergio & Sim-Sim
a
— Italy (1)
O. cristata Garside ex Perold
c
— Lesotho (1)
Ricciaceae
Riccia subgenus Ricciella
R. canaliculata Hoffm.
c
— UK (2)
R. cavernosa Hoffm. — Lesotho (2), UK (2)
R. crystallina L.
c
— Lesotho (3), UK (1)
R. fluitans L. — UK (4)
R. huebeneriana Lindb. — UK (1)
R. stricta (Lindb.) Perold
c
— Lesotho (2), Botswana (1)
Riccia subgenus Riccia
R. albolimbata S.W.Arnell
c
— Botswana (1)
R. beyrichiana Hampe ex Lehm. — UK (1)
R. crozalsii Levier
c
— Italy (1), UK (2)
R. glauca L. — UK (4)
R. montana Perold
c
— Lesotho (1)
R. nigrella DC.
c
— Italy (1), Lesotho (2), New Zealand (1), UK (2)
R. okahandjana S.W.Arnell
c
— Botswana (1)
R. sorocarpa Bisch. — UK (2)
R. subbifurca Croz.
c
— UK (3)
Ricciocarpus natans (L.) Corda — UK (2)
Jungermanniopsida, Metzgeriidae (simple thalloid liverworts):
Phyllothalliaceae
Phyllothallia nivicola A. E. Hodgs.
c
— Chile (1), New Zealand (1)
Fossombroniaceae
Austrofossombronia australis (Mitt.) R.M. Schust.
c
G New Zealand (1)
Fossombronia angulosa (Dicks.) Raddi G France (1), Italy (1), UK (3)
F. caespitiformis De Not. ex Rabenh.
c
G Italy (1)
F. echinata MacVicar
ac
G Italy (2)
F. pusilla (L.) Nees G UK (3)
F. maritima (Paton) Paton G UK (!)
F. wondraczeckii (Corda) Dum. ex Lindb. G UK (3)
Petalophyllum ralfsii (Wils.) Nees & Gottsche
a
G Italy (1), UK (3)
Allisoniaceae
Allisonia cockaynii (Steph.) R.M. Schust.
ac
G New Zealand (4)
Pelliaceae
Noteroclada confluens Tayl. ex Hook. & Wilson
a
G Chile (3), Venezuela (1)
Pellia endiviifolia (Dicks.) Dum.
a
G Italy (1), UK (4)
P. epiphylla ( L.) Corda
a
G UK (5), USA (2)
P. neesiana (Gottsche) Limpr. G UK (4)
Pallaviciniaceae
Greeneothallus gemmiparus Hassel
ac
G Chile (1)
Jensenia connivens (Colenso) Grolle
ac
G Venezuela (1)
J. wallichii Colenso
ac
G Venezuela (1)
Moerckia hibernica (Hook.) Gottsche
a
— (3) UK (2)
M. blyttii (Moerch) Brockm. G Switzerland (1), UK (4)
Pallavicinia connivens (Colenso) Steph.
ac
G New Zealand (2)
P. xiphoides (Hook.f. & Taylor) Trevis.
a
— (3) New Zealand (2)
P. tenuinervis (Hook.f. & Taylor) Trevis.
c
— New Zealand (2)
P. indica Schiffn.
c
— Malaysia (1)
P. lyellii (Hook.) Gray
a
— (2) UK (1), USA (1)
Podomitrium phyllanthus (Hook.) Mitt.
c
G New Zealand (1)
Symphyogyna brasiliensis Nees & Mont.
a
G South Africa (1), Venezuela (1)
S. brogniartii Mont.
a
G Venezuela (1)
S. hymenophyton (Hook.) Mont. & Nees
c
G New Zealand (4)
S. subsimplex Mitt.
c
G New Zealand (2)
S. undulata Colenso
c
G New Zealand (2)
Xenothallus vulcanicolus R.M. Schust.
ac
G New Zealand (1)
Hymenophytaceae
Hymenophyton flabellatum (Labill.) Dum.
a
G (1) New Zealand (4)
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

Immunocytochemistry—Epitopes associated with cell wall polysaccha-
rides and proteins were localized immunocytochemically in Marchantia
polymorpha subsp. montivagans and Conocephalum conicum. Colonized parts
of the thalli were cut into 0.5-mm-thick slices and fixed with 3% glutataldehyde
in 0.05 M PIPES buffer, pH 7.4 for 2 h at room temperature. After careful
rinsing in buffer, the samples were dehydrated in a step gradient of ethanol,
slowly infiltrated with LR White resin (Polysciences, Warrington, Pennsylva-
nia, USA), and cured at 608C for 24 h. The protocols followed for
immunohistochemistry and immunogold electron microscopy have been
described in detail in Ligrone et al. (2002). The antibodies tested, their
specificity, and source are listed in Table 4. For both light and electron
microscopy, controls were routinely made by omitting the incubation step with
the primary antibody and were always completely negative.
Resynthesis experiments—The apical parts of wild thalli of C. conicum,
about 2 mm long, were isolated and surface-sterilized with hypochlorite for 3
min, washed thoroughly in sterile distilled water, and placed on 0.25% phytagel
(Sigma) plates either lacking nutrients or containing one-fourth MS nutrient
solution (Murashige and Skoog, 1962). The plates were kept in a Sanyo MLR-
350 H growth chamber (Sanyo, Moriguchi City, Osaka, Japan)under a 12 h/12
h day/night photoperiod with a light irradiance of 50 Wm
2
and a 12/108C day/
TABLE 1. Continued.
Liverwort taxa Fungal status Geographic origin
Makinoaceae
Verdoornia succulenta R.M. Schust.
ac
B New Zealand (2)
Aneuraceae
Aneura lobata subsp. australis R.M. Schust.
ac
B New Zealand (4)
A. maxima Schiffn. (Steph.)
a
B (3) USA (1)
A. novaeguineensis Hewson
ac
B New Zealand (1)
A. pinguis (L.) Dum.
a
B UK (4)
A. pseudopinguis (Herzog) Pocs
ac
B Lesotho (2)
Cryptothallus mirabilis Malmb
a
B UK (4)
Riccardia intercellula E.A.Brown
c
B New Zealand (1)
R. pennata E.A.Brown
c
B New Zealand (1)
R. chamedryfolia (With.) Grolle — (3) UK (4)
R. cochleata (Hook.f. & Taylor) Kuntze
c
— New Zealand (1)
R. eriocaula (Hook.) Besch. & C.Massal.
c
— New Zealand (2)
R. incurvata Lindb. — (4) UK (5)
R. latifrons (Lindb.) Lindb. — (4) UK (3)
R. multifida (L.) Gray — (3) UK (3)
R. palmata (Hedw.) Carruth. — (3) UK (3)
Metzgeriaceae
Apometzgeria pubescens (Schrank) Kuwah. — UK (2)
Metzgeria conjugata Lindb. — UK (3)
M. decipiens (Massal.) Schiffn. & Gotts. — Chile (3)
M. temperata Kuwah.
a
—UK(2)
M. furcata (L.) Dum
a
— UK (4)
M. fruticulosa (Dicks.) A. Evans — UK (2)
Pleuroziaceae
Pleurozia purpurea Lindb. — UK (3)
P. gigantea (Web.) Lindb.
c
— Malaysia (1)
Notes: G, glomeromycotean endophytes; B, basidiomycotean endophytes; —, fungal endophytes absent. Numbers in parentheses after each country of
origin refer to the number of voucher specimens examined (see Appendix).
a
Liverwort taxa examined by electron microscopy in this and our previous studies.
b
Herbarium specimens only.
c
Liverwort taxa not included in the previous survey by Nebel et al. (2004); under column ‘‘Fungus status’’: (1) taxon reported by Nebel et al. (2004) as
nonmycorrhizal or as associated with (2) glomeromycotean fungus, (3) unidentified fungus, or (4) basidiomycetous fungus. Unless indicated otherwise, the
fungal status agrees with that reported by Nebel et al. (2004) for the same species. No glomeromycotean associations were found in the Jungermanniidae
(leafy liverworts).
TABLE 2. Restriction profiles of fungal small-subunit rDNA 550-bp
amplicons from fungus-associated liverworts.
Restriction enzyme Restriction profiles (bp)
Hinf I H1 (383, 120, 5)
H2 (244, 188, 90, 25)
H3 (383, 141, 25)
H4 (334, 190, 25)
H5 (334, 141, 49, 25)
H6 (278, 244, 25)
Hsp92II S1 (249, 148, 90, 23)
S2 (291, 163, 93)
S3 (291, 258)
S4 (291, 142, 116)
TABLE 3. Combinations of restriction profiles with Hinf I (H1-H6) and
Hsp92II (S1-S4), and GenBank sequence codes (in parentheses) of
fungal small-subunit rDNA 550-bp amplicons from fungus-associated
liverworts.
Haplomitrium chilensis H2S2 (AM412526, AM412528)
Conocephalum conicum H4S2 (AM412536, AM412537, AM412538)
H4S3 (AM412539)
H3S4 (AM412540, AM412541)
Monoclea gottschei H1S1 (AM412542, AM412543, AM412544,
AM412545)
Fossombronia echinata H5S3 (AM 412532, AM412535)
H2S3 (AM 412534)
H2S2 (AM 412533)
Pellia endiviifolia H3S3 (AM 412529)
H6S2 (AM 412530)
AMERICAN JOURNAL OF BOTANY [Vol. 94

night temperature regime. After 3 mo in culture, plates containing thalli about
10 mm long were inoculated with glomeromycotean spores that had been
previously surface-sterilized with 3% chloramine T and 0.03% streptomycin for
5 min. Four different glomeromycotean species were tested: Gigaspora rosea
Nicolson & Schenck (BEG9) and Gigaspora margarita Becker & Hall
(BEG34), maintained at the Istituto per la Protezione delle Piante, Torino, Italy,
and Glomus mosseae (Nicol. & Gerd.) Gerd. & Trappe (BEG12) and Glomus
clarum Nicolson & Schenck (BEG142), kindly supplied by Dr. Vivienne
Gianinazzi-Pearson (INRA, Dijon, France). The cultures were examined at
intervals with a dissection microscope. Putative associations, identified from
fungal colonization of rhizoids and of the internal tissue of the thalli, were
processed for light and electron microscopy as described.
RESULTS
Molecular identification of fungal endophytes—PCR
amplification of DNA from fungus-colonized liverwort tissue
with the NS31 and AM1 primers produced a DNA fragment of
about 550 bp. RFLP analysis of this fragment with the
restriction enzymes Hinf I and Hsp92II produced six and four
different RFLP types, respectively (Table 2). One to three
different restriction patterns were obtained from each liverwort
species, and for each pattern one to four amplicons were
sequenced (Table 3). When the sequences were aligned with a
data set from GenBank, they all clustered within the
Glomeromycota with high bootstrap support, producing a tree
topology coherent with those from Schu¨ ßler et al. (2001) (Fig.
1). The sequence from Monoclea was closely related with
Acaulospora, while the remaining sequences were related with
the Glomus Group A, in part clustering within this group and in
part forming a sister clade to it (Fig. 1).
The taxonomic distribution of glomeromycotean associa-
tions in liverworts—The cytology of the GAs for the fungal
endophytes that were identified by molecular analysis (see next
section) was used as a reference for morphological identifica-
tion of fungal endophytes in the other taxa listed in Table 1.
Diagnostic features for GAs were absence of visible pathogenic
symptoms in host plants, intracellular colonization by aseptate
hyphae, fungal colonization restricted to specific tissue areas in
the gametophyte (see next section) and absent from the
sporophyte, fungal entry via the rhizoids (except in the
Haplomitriopsida, the development of intracellular arbuscule-
like structures, the development of fungal vesicles, and
endobacteria occurring in fungal hyphae. Septate fungi were
identified by electron microscopy as basidiomycetes or
ascomycetes from the presence of dolipores or simple septa,
respectively.
While in the majority of GA-forming taxa the fungal
association was consistently present regardless of the collecting
site or season, in a few species the degree of colonization was
highly variable from plant to plant even within the same
sample. For example, populations of Monoclea forsteri, M.
gottschei,Conocephalum conicum, Lunularia cruciata, Du-
mortiera hirsuta, and Noteroclada confluens growing either in
very wet or epilithic habitats were more variable than were
populations growing on soil. In Marchantia polymorpha, the
fungus was present in the subspecies montivagans, a perennial
taxon growing in natural habitats, but was absent from the two
pioneer subspecies, polymorpha and ruderalis, that colonize
ephemeral and usually nutrient-rich habitats. Also lacking
endophytes were the linear branches with very few rhizoids in
Pellia spp. from wet habitats and the furcate caducous rhizoid-
free branches of P. endiviifolia that proliferate in the autumn
and early winter (Paton, 1999). For each liverwort taxon
examined, GAs were reported as present when consistently
found in at least a part of the specimens examined, provided
that the morphological criteria detailed previously were
satisfied. Our reports refer to the potential ability, or apparent
inability (the latter amenable to confutation by examination of
further samples), of certain taxa to establish this type of
symbiosis, with no assumption of ecological relevance.
Moreover, no attempt was made in this study to quantitatively
evaluate the occurrence of the fungi.
Based on the guidelines described, GAs were found to be
widespread in a large liverwort assemblage encompassing the
Haplomitriopsida, Marchantiopsida, and part of the Metzger-
iidae within the Jungermanniopsida (Table 1). Within the
Marchantiopsida, fungal endophytes were consistently absent
in a minority of taxa, notably the Blasiales, Sphaerocarpales,
and within the Marchantiales in the families Wiesnerellaceae,
Monoseleniaceae, Exormothecaceae, Cyathodiaceae, Mono-
carpaceae, Oxymitraceae, and Ricciaceae. Within the Metzger-
TABLE 4. Monoclonal antibodies utilized for immunocytochemical characterization of cell walls in liverworts.
Antibody Antigen(s)/epitope Reference/source
Anti-callose Callose/penta- to hexa-(1!3)-b-glucan Meikle et al., 1991/Biosupplies Ltd, Melbourne, Australia
CCRC-M1 Xyloglucan, rhamnogalacturonan-I/terminal (1!2)-a-linked
fucosyl-containing side group
Puhlman et al., 1994/M. Hahn, Complex Carbohydrate Research Center,
University of Georgia, USA
CCRC-M2 Rhamnogalacturonan-I/unknown Puhlman et al., 1994/M. Hahn, Complex Carbohydrate Research Center,
University of Georgia, USA
CCRC-M7 Arabinogalactan-proteins, rhamnogalacturonan-I/
arabinosilated (1!6)-b-galactan
Puhlman et al., 1994; Steffan et al., 1995/M. Hahn, Complex Carbohydrate
Research Center, University of Georgia, USA
LM1 Hydroxyproline-rich glycoproteins/undefined Smallwood et al., 1996/J.P. Knox, Centre for Plant Sciences,
University of Leeds, UK
LM2 Arabinogalactan-proteins/undefined Smallwood et al., 1995/J.P. Knox, Centre for Plant Sciences,
University of Leeds, UK
LM5 Galactan, rhamnogalacturonan-I/tetra (1!4)-b-galactan Jones et al., 1997/J.P. Knox, Centre for Plant Sciences, University of Leeds, UK
LM6 Arabinan, rhamnogalacturonan-I/penta(1!5)-a-arabinan Willats et al., 1998/J.P. Knox, Centre for Plant Sciences, University of Leeds, UK
LM7 Homogalacturonan/partially methyl esterified Willats et al., 2001/J.P. Knox, Centre for Plant Sciences, University of Leeds, UK
JIM5 Homogalacturonan/low- or unmethyl-esterified Willats et al., 2000/J.P. Knox, Centre for Plant Sciences, University of Leeds, UK
JIM7 Homogalacturonan/partially methyl-esterified Willats et al., 2000/J.P. Knox, Centre for Plant Sciences, University of Leeds, UK
JIM11 Hydroxyproline-rich glycoproteins/undefined Smallwood et al., 1994/J.P. Knox, Centre for Plant Sciences,
University of Leeds, UK
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

Fig. 1. Phylogenetic tree from 550-bp fungal small-subunit rDNA data sets from five fungus-associated liverworts. Bootstraps values above 75% are
reported at the nodes. The sequences from the liverworts are in boldface type. The tree was rooted with Mortierella polycephala (Zygomycota), Ustilago
hordei (Basidiomycota), and Neurospora crassa (Ascomycota). The bar at the base of the diagram is a measure of phylogenetic distance.
AMERICAN JOURNAL OF BOTANY [Vol. 94

iidae clade I (Davis, 2004), GAs were common, with the
exception of a few isolated species in the Pallaviciniaceae,
while the remaining taxa traditionally included in the
Metzgeriidae and grouped in the Metzgeriidae clade II by
Davis (2004) were either fungal free (Pleuroziaceae and
Metzgeriaceae) or associated with basidiomycetes (Aneuraceae
and Verdoornia). In no case has a putative GA been detected in
the Jungermanniidae (leafy liverworts) (Duckett et al., 2006b;
J. G. Duckett, unpublished data).
Cytology of glomeromycotean associations in liverworts—
GAs in Haplomitrium and Treubia have been described in
detail in previous papers (Carafa et al., 2003; Duckett et al.,
2006a) and will not be considered in this section. In the
Marchantiopsida and Metzgeriidae, including the species
investigated by molecular techniques, glomeromycotean colo-
nizations were typically localized in the rhizoids and the
internal parenchyma along the midrib of the thallus (Fig. 2A,
B). The meristematic regions up to 2–3 mm behind the apices,
the sex organs, and the sporophytes, including the placental
area associated with the foot, were never colonized. The oil-
body idioblasts in marchantialean liverworts remained fungal
free even when surrounded by colonized cells. Also fungus-
free was the strand of hyaline cells occupying the lower part of
Fig. 2. Light micrographs of glomeromycotean associations in the gametophyte of (A) Asterella wilmsii, (B) Monoclea forsteri, (C) Marchantia
polymorpha subsp. montivagans, and (D) Symphyogyna brasiliensis, showing colonized areas (F) of the internal parenchyma. Scale bars: A–D, 100 lm.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

the midrib in certain marchantialean taxa such as Conocepha-
lum and Marchantia (Fig. 2C). In many members of the
Pallaviciniaceae, GAs were restricted to subterranean stolons
lacking a laminar margin (Fig. 2D). Fungi were also rare or
absent in the lipid-laden regions of the perennating tubers of
Petalophyllum ralfsiii and Fossombronia maritima.
Unlike the Haplomitriopsida (Carafa et al., 2003; Duckett et
al., 2006a), direct fungal penetration through epidermal cells
was never observed in the Marchantiopsida or Metzgeriidae,
indicating that here the rhizoids are the only access to the
fungus. The fungus penetrated the rhizoids at any point and
formed large intracellular hyphae running in both directions
(Fig. 3A, B). Of the two types of rhizoids present in many
marchantialean liverworts, i.e., living smooth rhizoids and
tuberculate rhizoids that undergo cytoplasmic lysis at maturity,
only the former were found to be primarily colonized by
glomeromycotean fungi. From rhizoids, the hyphae entered the
parenchyma cells above the lower epidermis of the thallus (Fig.
3C, D). The colonization was entirely intracellular and closely
resembled Paris-type arbuscular mycorrhizas (Smith and
Smith, 1997), with large colonizing hyphae spreading from
cell to cell and intercalary formation of arbuscule-like
structures from shorter lateral branches, or trunk hyphae, with
determinate growth (Fig. 3E, F). In taxa with large thalli, such
as Conocephalum conicum or Marchantia polymorpha, the
colonizing hyphae often grew longitudinally along the midrib
of the thallus, probably extending the colonization to a
significant distance from the entry point (Fig. 4A). In most
other taxa, however, the hyphae had no particular orientation in
the thallus parenchyma. A particularly well-differentiated
Fig. 3. (A–C) Light micrographs and (D–E) scanning electron micrographs of glomeromycotean associations in liverworts. (A) Fungus-colonized
rhizoid of Conocephalum conicum; the arrow points to the penetration site of a fungal hypha. (B) Rhizoid of Marchantia polymorpha subsp. montivagans
containing fungal hyphae and vesicles (arrows). (C) Rhizoid base with fungal hypae (arrow) in Monoclea forsteri. (D) Fungal hyphae (arrow) passing from
the rhizoid base (R) to adjacent parenchyma cells in Preissia quadrata. (E) Colonizing hypha crossing host cell walls (arrows) and (F) arbuscule in the
thallus parenchyma of Fossombronia echinata. Scale bars: A–D, 20 lm; E, F, 10 lm.
AMERICAN JOURNAL OF BOTANY [Vol. 94

association was found in the genus Marchantia. Here the
fungus colonized a region overarching the midrib hyaline
strand and consisting of a lower area containing coils and
vesicles and an upper area containing arbuscules (Fig. 4B–D).
Cell wall autofluorescence, a normal feature of fungus-free
parenchyma cells in the thallus of Conocephalum and other
marchantialean liverworts, was no longer visible in colonized
cells (Fig. 5A). Autofluorescence disappeared first from the
cell walls crossed by the fungus (Fig. 5B, C).
Intracellular vesicles, both in rhizoids and parenchyma cells,
developed in most of the taxa examined (Figs. 3B and 4C).
During fungal penetration, the host cell wall underwent local
lysis, while the host plasmalemma invaginated to form a
perifungal membrane that surrounded the intracellular fungus
and separated it from the host cytoplasm. An interfacial matrix
of fibrillar material was deposited in the space between the
hyphae and perifungal membrane, with a thickness decreasing
from 0.5–1.0 lm at entry/exit points, where it formed a
conspicuous collar around the fungus (Fig. 5D), to 50 nm or
less in fine arbuscular hyphae (Fig. 6E). The colonizing hyphae
were 3–6 lm in diameter, rarely less, and had relatively thick
walls (100–200 nm), sometimes with a layered structure (Fig.
6A); the fungal cell walls retained the same thickness or
became slightly thinner in larger trunk hyphae, but they thinned
to about 30 nm or less in terminal arbuscular hyphae (Fig. 6E).
The colonizing hyphae contained numerous vacuoles with
electron-transparent contents and irregular shapes; scattered in
the cytoplasm were several nuclei, mitochondria, and mem-
Fig. 4. Light micrographs of glomeromycotean associations in liverworts. (A) Detail of the thallus parenchyma in Conocephalum conicum showing
large colonizing hyphae (arrows) growing along the longitudinal axis of the thallus. (B) Colonized area in the thallus midrib of Marchantia polymorpha
subsp. montivagans consisting of a lower region with fungal coils and vesicles (C) and an upper region with arbuscules (A). (C, D) Details of the (C) lower
and (D) upper region; a vesicle (V) and a fungus-free oil-body idioblast (OB) are visible in (C) and a large colonizing hypha (CH ) and arbuscules (A)in
(D). Scale bars: A, 40 lm; B, 100 lm; C, D, 20 lm.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

brane-bound spheroidal bodies of electron-opaque material
(Figs. 5D and 6A, B). The trunk hyphae of the arbuscules
appeared similar to colonizing hyphae, but often they could be
distinguished because their cytoplasm was filled with minute
vacuoles (Fig. 5E). With very few exceptions (e.g., the fungal
endophytes in Haplomitrium gibbsiae and Pellia epiphylla),
endocellular bacteria were present in both colonizing and trunk
hyphae. These appeared as cocci about 0.3–0.5 lm in diameter,
sometimes of more irregular shape, with an electron-opaque
cell wall of the gram-positive type (i.e., relatively thick and
lacking an outer membrane) and no bounding fungal
membrane. Division of bacterial endophytes by a central
constriction was observed frequently (Fig. 6C). The terminal
arbuscular hyphae typically were less than 1 lm in diameter
and contained no nuclei nor endobacteria (Fig. 6E). As reported
for Treubia (Duckett et al., 2006a), the fungal endophyte in
Petalophyllum did not form typical arbuscules but only coiled
hyphae of relatively uniform diameter (Fig. 6D). The
colonizing hyphae in Petalophyllum did not exceed 3 lmin
diameter, and the thinner intracellular hyphae were rarely less
than 1 lm.
With fungal colonization, the host cells underwent pro-
nounced morphological changes that were remarkably uniform
in all taxa investigated. These included proliferation of the
Fig. 5. (A) Cell-wall autofluorescence in the thallus parenchyma of Conocephalum conicum; note the absence of fluorescence in the fungus-colonized
area (F). The red fluorescence is from chloroplasts in the adjacent chlorenchyma. (B) Bright-field micrographs of colonizing hypha spreading in the thallus
parenchyma of C. conicum; the arrows point to host cell walls crossed by the fungus. (C) Fluorescence micrographs of the same area showing that
autofluorescence first disappears in cell walls crossed by the fungus (arrows). (D, E) Transmission electron micrograph of fungus-colonized thallus
parenchyma cells in Marchantia polymorpha subsp. montivagans. (D) Detail of colonizing hypha crossing a host cell wall; the arrows point to the collars
of interfacial material; fungal nucleus (N); perifungal membrane continuous with host plasmalemma (arrowheads). (E) Detail of an arbuscule-containing
cell, showing the host nucleus (HN ), a large trunk hypha (TH ), and numerous arbuscular hyphae (AH ) surrounded by the host cytoplasm. Scale bars: A,
100 lm; B, C, 40 lm; D, 1 lm; E, 2 lm.
AMERICAN JOURNAL OF BOTANY [Vol. 94

Fig. 6. Transmission electron micrographs of glomeromycotean associations in liverworts. (A) Transverse section of a colonizing hypha in Pellia
epiphylla; the arrow points to the thick multilayered wall. (B) Detail of colonizing hypha in P. epiphylla showing membrane-bound, electron-opaque
bodies (arrows). (C) Bacterial endophyte with central constriction in a trunk hypha in Marchantia paleacea; note the gram-positive type bacterial wall and
the absence of a bounding fungal membrane. (D) Fungus-colonized parenchyma cell in Petalophyllum ralfsii; note absence of a typical arbuscule. (E)
Detail of an arbuscule-containing cell in M. paleacea; the arbuscular hyphae (AH ) establish intimate spatial relationships with host organelles including
mitochondria (M), microbodies (Mb), and plastids (P). Note the absence of starch in plastids. A dictyosome (G) and several profiles of endoplasmic
reticulum are also visible in the host cytoplasm. Scale bars: A, 1 lm; B, 0.2 lm; C, 0.1 lm; D, 2 lm; E, 0.5 lm.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

Fig. 7. Transmission electron micrographs of glomeromycotean associations in liverworts. (A) Oil bodies (OB) in fungus-free and (B) colonized
parenchyma cell in Petalophyllum ralfsii, showing differences in the appearance of the matrix and lipid components; note the starch-filled plastids (P)in
the fungus-free cell. (C) Intravacuolar clump of collapsed hyphae in Hymenophyton flabellatum. (D) Degenerated hyphae (F)inP. ralfsii; the arrow points
to the ghost of a crystal. (E) Fungal vesicle at an early stage of development in Marchantia foliacea; note the thin wall (arrow) and numerous nuclei (N)
scattered in the cytoplasm. (F) Fungal vesicle at a more advanced stage of development in Pellia epiphylla; the fungal wall (arrow) has become much
thicker and the cytoplasm is packed with lipid (L). Scale bars: A, C, 1 lm; B, D, 0.5 lm; E, F, 2 lm.
AMERICAN JOURNAL OF BOTANY [Vol. 94

cytoplasm and organelles, replacement of the large central
vacuole typical of fungus-free cells with numerous smaller
vacuoles separated by cytoplasmic strands, migration of the
nucleus from a peripheral to an internal position, and
disappearance or strong reduction of starch in plastids. The
nucleus and plastids often became pleomorphic. The fungus
established intimate spatial relationships with the nucleus,
plastids, mitochondria, and other host organelles (Figs. 5E and
6E). In the Metzgeriopsida (in which all somatic cells typically
contain oil bodies) the density of the matrix and the abundance
of lipid droplets in the oil bodies markedly declined in
colonized cells (Fig. 7A, B).
At a more advanced stage of colonization, the arbuscules
degenerated, forming one to several clumps of collapsed
hyphae. The larger hyphae usually survived the arbuscules and
could occasionally give rise to a second colonization cycle,
inferred from the presence of a healthy arbuscular system along
with clumps of collapsed hyphae in the same cells. When all
the intracellular fungus was dead, the host cells resumed their
precolonization cytological organization, the only sign of past
colonization being intravacuolar clumps of fungal wall remains
(Fig. 7C). A different pattern of fungal degeneration was
observed in Petalophyllum. Here the hyphae underwent cell
wall dissolution and cytoplasmic lysis, producing masses of
amorphous material in which no fungal walls were discernible
(Fig. 7D). Common in degenerated hyphae in Petalophyllum
were ghosts of crystals, probably calcium oxalate, that
dissolved during fixation (Fig. 7D).
Vesicles developed by terminal swelling of colonizing
hyphae or of lateral branches. Initial vesicle development was
characterized by nuclear and organelle proliferation (Fig. 7E);
at later stages the vesicles accumulated abundant lipid reserves
and their cell walls thickened conspicuously (Fig. 7F).
Immunocytochemistry—The immunocytochemical tests in
C. conicum and M. polymorpha produced very similar results
(Table 5). The antibody against (1!3)-b–glucan strongly
labeled the host wall material associated with plasmodesmata
(Fig. 8A), while no labeling was observed at the level of fungal
penetration nor in the interfacial material covering the
intracellular hyphae. The same antibody also labeled the wall
of hyphae external to the thallus (Fig. 8B) but not of
intracellular hyphae; in the latter some labeling was observed
only within the vacuoles (Fig. 8C). JIM5 and JIM7, two
antibodies against homogalacturonan, and JIM11, which
recognizes an epitope associated with hydroxyprolyne-rich
proteins, labeled the liverwort cell walls throughout (except the
cell corners) as well as the interfacial material associated with
the intracellular fungus (Fig. 8D–F).
Perhaps the most interesting results were those obtained with
CCRC-M1, a monoclonal antibody that recognizes fucosylated
side groups associated with xyloglucan and rhamnogalactur-
onan I (Puhlman et al., 1994). This antibody produced very
little labeling of the cell walls in fungus-free cells, including
meristematic cells. In contrast, the same antibody strongly
labeled the interfacial material associated with the intracellular
fungus (Fig. 9A, B). Moreover, starting from the penetration
site, colonized cells deposited a new wall layer that was
continuous with the interfacial material and was also heavily
labeled by CCRC-M1 (Fig. 9C).
In vitro synthesis of glomeromycotean associations—The
four fungal isolates tested were all able to colonize the roots of
the higher plant Trifolium repens L., producing typical AMs. In
contrast, successful colonization of the host liverwort (C.
conicum) was obtained only with spores of Glomus mosseae
and only in about 10% of the plants inoculated. In the other
cases, the fungal spores either failed to germinate (Glomus
clarum) or produced germlings that stopped growing and died
(Gigaspora rosea and G. margarita, and some G. mosseae
spores).
The colonized plants were maintained in culture for several
months with no adverse symptoms, although fungal coloniza-
tion did not appreciably enhance their growth relative to the
TABLE 5. Immunogold labeling in mature thallus parenchyma of the liverworts Conocephalum conicum and Marchantia polymorpha subsp. montivagans
(Marchantiopsida).
Antibody
C. conicum M. polymorpha
Liverwort cells Fungal hyphae Liverwort cells Fungal hyphae
Anti-b-glucan þþþ Plasmodesmatal collars þþ Cell walls in
external hyphae
þþþ Plasmodesmatal collars þþ Cell walls in
external hyphae
Rest of cell walls Cell walls in
internal hyphae
Rest of cell walls Cell walls in
internal hyphae
þVacuoles þVacuoles
CCRC-M1 6Cell walls in fungus-free cells 6Cell walls in fungus-free cells
þþ Cell walls in colonized cells
and interfacial material
 þþþ Cell walls in colonized cells
and interfacial material

CCRC-M2   
CCRC-M7   
LM1   
LM2   
LM5   
LM6   
LM7 þ þ 
JIM5 þþþ  þþþ 
JIM7 þþþ  þþþ 
JIM11 þþþ  þþþ 
Notes: Relative intensity of labeling: þþþ very strong, þþ strong, þweak, 6very weak and uneven, – absent. Positive reports with no further
information indicate nonspecific labeling of all host cell walls and of the interfacial material.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

controls. The synthesized association developed through the
same steps as observed in wild plants; the fungus first entered
the rhizoids and subsequently colonized the thallus parenchy-
ma by growing from cell to cell and produced intracellular
arbuscules (Fig. 10A). Apart from being more highly
vacuolated, a likely consequence of growth in a water-saturated
environment, colonized parenchyma cells in the synthesized
association were morphologically indistinguishable from their
wild counterparts (Fig. 10B).
DISCUSSION
The nature of fungal endophytes—Of the five liverwort
species selected for molecular analysis, three were from Europe
(Conocephalum, Fossombronia, and Pellia), one was from
New Zealand (Monoclea), and one was from South America
(Haplomitrium). Molecular analysis demonstrates that these
species all contain fungal endophytes that cluster with the
Glomeromycota and are related either to Glomus Group A
(Schwarzott et al., 2001) or, in the case of Monoclea,to
Fig. 8. Immunocytochemistry of glomeromycotean associations in liverworts. (A–C) Localization of (1!3)-b–glucan epitopes in Conocephalum
conicum. (A) Labeling of the host cell wall around the plasmodesmata (arrows), indicating the presence of callose. (B) Labeling of the fungal wall in
external hyphae (arrows). (C) Detail of a colonizing hypha at the penetration point; no labeling is visible in the fungal wall (FW) or in the interfacial
material (IM ); some labeling is visible in fungal vacuoles (FV ); host cell wall (HW ). (D–F) Details of colonizing hyphae (F) and host cell wall at
penetration points in Marchantia polymorpha subsp. montivagans, showing labeling with (D) JIM7, (E) JIM5, and (F) JIM11; these antibodies labeled
both the host walls (HW ) and interfacial material (IM ). Scale bars: A–F, 0.3 lm.
AMERICAN JOURNAL OF BOTANY [Vol. 94

Acaulospora. Both glomeromycotean lineages form arbuscular
mycorrizas in tracheophytes (Smith and Read, 1997; Peterson
et al., 2004). The results are consistent with a former study that
demonstrated the presence of glomeromycotean endophytes
related to Glomus Group A in populations of Marchantia
foliacea in New Zealand (Russell and Bulman, 2005). In line
with molecular analysis, our resynthesis experiments showed
that G. mosseae, a glomeromycotean fungus that nests within
the Glomus Group A (Fig. 1), was able to colonize axenic thalli
of C. conicum and to establish an endophytic association
closely similar to that observed in the wild. A similar result was
obtained in a cross-colonization experiment with the simple
thalloid liverwort Pellia epiphylla and an unidentified
glomeromycotean fungus associated with the higher plant
Plantago lanceolata (Read et al., 2000). The low frequency of
colonization observed in Conocephalum after inoculation with
G. mosseae and the total failure with the other glomeromyco-
tean isolates tested in the present study may reflect low
compatibility and/or an inhibitory effect of growth conditions
on the liverwort ability to elicit fungal development. In line
with the first possibility is the repeated occurrence of the same
fungal phylotypes in populations of M. paleacea from different
sites (Russell and Bulman, 2005). Although too few taxa have
been studied to support any general conclusion, the data
suggest a degree of specificity between liverworts and Glomus
Group A that contrasts with the large spectrum of glomero-
mycotean associates in tracheophytes (Peterson et al., 2004).
The taxonomic distribution and origins of GAs in
liverworts—The application of diagnostic criteria inferred
Fig. 9. Immunogold labeling with CCRC-M1 monoclonal antibody in
glomeromycotean associations in liverworts. (A) Colonizing hypha (F)
crossing a host cell wall (HW )inMarchantia polymorpha subsp.
montivagans; the antibody labeled the interfacial material covering the
fungus (arrows). (B) Detail of (A), showing heavy labeling of interfacial
material at the fungal entry point (arrow). (C) Detail of a cell wall at the
interface between a fungus-colonized (CC ) and a fungus-free (UC ) host
cell; a heavily labeled cell wall layer (bracket) is visible on the side
towards the colonized cell while no labeling is visible on the other side of
the cell wall. Scale bars: A–C, 0.3 lm.
Fig. 10. Resynthesis of a glomeromycotean association from spores of
Glomus mosseae and axenic thalli of Conocephalum conicum. (A) Hand
section of a thallus stained with aniline blue, showing fungus-colonized
rhizoids (arrows) and parenchyma cells (IP). (B) Transmission electron
micrographs of colonized parenchyma cell, showing profiles of colonizing
hypha (CH ), trunk hyphae (TH ), and arbuscular hyphae (AH ). Scale bars:
A, 40 lm; B, 3 lm.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

from the cytological analysis of the liverwort species with
fungal endophytes that we identified by molecular techniques
has provided more solid support for the morphological
identification of GAs in other taxa. With information on 67
species with a previously unknown fungal status and the
reexamination of 64 species already included in the list by
Nemec et al. (2004), our survey confirms GAs as a general
feature of a large liverwort assemblage encompassing the
Haplomitriopsida, most of the Marchantiopsida, and part of the
Metzgeriidae (the simple thalloid clade I according to Davis,
2004). With reference to the topology of the phyletic tree of
liverworts produced by cladistic analysis (Forrest and Crandall-
Stotler, 2004, 2005; Heinrichs et al., 2005; Forrest et al., 2006)
and shown in a simplified version in Fig. 11, this taxonomic
distribution strongly suggests that the symbiotic association
with glomeromycotean endophytes is a plesiomorphy in
liverworts. Accordingly, the consistent absence of GAs in
certain taxa, both basal (Blasiales and Sphaerocarpales) and
derived (Ricciaceae, the simple thalloid clade II and the whole
clade of leafy liverworts) should be interpreted as the result of
multiple independent losses. However, the apparent liverwort
tendency to associate predominantly with fungi related to the
Glomus Group A is consistent with host shifting of symbionts
from tracheophytes to liverworts (Selosse, 2005). The latter
hypothesis might explain in terms of multiple acquisitions, at
least in part, the scattered distribution of GAs in liverworts.
Discrepancies between the present study and the survey by
Nebel et al. (2004) relative to certain taxa, in particular
Corsinia coriandrina and Hymenophyton flabellatum (Table
1), may reflect intraspecific ecological variability. Further
investigation is needed to ascertain whether the absence of
fungal endophytes in several isolated species within mycorrh-
ized families, such as Cryptomitrium oreoides in the
Aytoniaceae or several species of Pallavicinia in the
Pallaviciniaceae, are further instances of multiple evolutionary
loss/acquisition or of ecological variability as noted in
Conocephalum, Lunularia, Pellia,Noteroclada, Dumortiera,
and Monoclea. As in vascular plants, many of the liverworts
that lack GAs grow in very wet habitats. Paradoxically,
however, absence is equally common in liverwort taxa growing
in places subjected to intense seasonal desiccation. The absence
of GAs from the two Marchantia polymorpha subspecies
growing in nutrient-rich habitats (polymorpha and ruderalis)is
not unexpected and suggests that shifting from the mycorrhizal
to nonmycorrhizal status in liverworts is relatively easy in
evolutionary terms.
Our survey confirms the absence of GAs in the Pleuro-
ziaceae and Metzgeriaceae, and we report the presence of
basidiomycetous endophytes not only in the Aneuraceae but
also in Verdoornia, a taxon traditionally placed in the distantly
related family Makinoaceae (Crandall-Stotler and Stotler,
2000). In molecular phylogenies these four groups form a
single clade (simple thalloid II, Fig. 11) with a sister
relationship to the leafy liverworts (Davis, 2004; Forrest and
Crandall-Stotler, 2004; Heinrichs et al., 2005). More detailed
analysis is now needed to ascertain possible affinities of the
basidiomycete associations in Verdoornia and in the Aneur-
aceae (Kottke et al., 2003) and thereby to gain insight into the
evolution of these associations following the postulated loss of
GAs in the common ancestor to the simple thalloid II/leafy
liverwort lineage (Kottke and Nebel, 2005).
Morphological and cellular aspects—GAs in the Marchan-
tiopsida and Metzgeriidae are remarkably uniform in develop-
ment and morphology. In contrast, GAs in the
Haplomitriopsida have several unique features including the
colonization of epidermal cells in Haplomitrium, the coloni-
zation of intercellular spaces in Treubia, and the development
of thin-walled hyphal swellings in both genera (Carafa et al.,
2003; Duckett et al., 2006a). Because molecular analysis has
shown that the fungal endophyte of H. chilensis clusters with
the endophytes from marchantialean and metzgerialalean
liverworts, the distinctive morphology of GAs in the
Haplomitriopsida appears to depend on control by the host
rather than the fungus.
The results of immunocytochemical analysis of GAs in
Conocephalum and Marchantia indicate a level of functional
interaction between the symbionts comparable to that in AMs.
No callose deposition was observed in colonized cells at the
points of fungal entry nor at the host/fungus interface. Callose
deposition has been implicated in numerous studies as a
resistance response to attack by pathogens (Rodriguez-Galvez
and Mendgen, 1995; Enkerli et al., 1997), while higher plants
produce little or no callose in reacting to AM fungi (Balestrini
et al., 1994; Gianinazzi-Pearson et al., 1996). Also significant
is the observation that the antibody against (1!3)-b-glucan
labels the cells walls of external hyphae but not those of
intracellular hyphae, suggesting that the association with the
host liverwort inhibits the synthesis of this polysaccharide in
the fungus. A decrease in cell wall labeling by antibodies
against (1!3)-b-glucan in AM fungi has been interpreted as a
sign of structural simplification of the fungal wall accompa-
Fig. 11. Phyletic tree of the liverworts and their position relative to the
rest of the embryophytes. The asterisks indicate the clades that form
endophytic associations with glomeromycetes. For further details about
liverwort and embryophyte phylogeny, see Dombrovska and Qiu (2004);
Forrest and Crandall-Stotler (2004, 2005); Forrest et al. (2006); Groth-
Malonek et al. (2005); Heinrichs et al. (2005); Qiu et al. (2006).
AMERICAN JOURNAL OF BOTANY [Vol. 94

nying the development of the intraradical phase (Lemoine et
al., 1995).
The cell walls in the thallus parenchyma of Marchantia and
Conocephalum were strongly labeled by antibodies against
homogalacturonans with different degrees of methyl esterifi-
cation (JIM5 and JIM7) and by an antibody that recognizes an
epitope associated with hydroxyprolyne-rich proteins (JIM11).
Both groups of compounds are widespread components of cell
walls in plants but are not known in fungal walls. Therefore,
the presence of the same epitopes in the interfacial matrix
ensheathing the intracellular mycobiont indicates that, as in
AMs (Balestrini et al., 1996; Harrison, 1997; Balestrini and
Bonfante, 2005), this material is of host origin and that the host
cells colonized by the fungus maintain the ability to synthesize
and secrete cell wall material. The results obtained with CCRC-
M1 demonstrate that the fungal colonization elicits the
synthesis of cell wall polysaccharide(s) that are scarcely
present in fungus-free thallus parenchyma cells. The suppres-
sion of autofluorescence in colonized cells also indicates
changes in cell wall composition consequent to fungal
colonization. No change in the expression of the CCRC-M1
epitope like that observed in this study has been reported in
other glomeromycotean associations. Immunogold labeling of
AMs in higher plants with CCRC-M1 and CCRC-M7 showed
that, although the tissue distribution of the epitopes of these
two antibodies varied according to the plant species, the
interfacial matrix invariably had the same labeling pattern as
that found in host cell walls before fungal colonization
(Balestrini et al., 1996). In contrast, fungal colonization in
cucumber AMs elicited the expression of two different
expansin proteins, one localized in the host cell walls and the
other in the interfacial matrix (Balestrini et al., 2005).
Endocellular bacteria are common in glomeromycotean
fungi forming AMs in higher plants. Originally reported as
‘‘bacterium-like organelles,’’ the glomeromycotean endobac-
teria were first studied by Macdonald et al. (1982), who
described three different types, either free in fungal cytoplasm
or enclosed in fungal membrane. Membrane-bound, rod-
shaped endobacteria in the glomeromycotean family Giga-
sporaceae have been identified as gram-negative b-proteobac-
teria related to the genus Burkholderia (Bianciotto et al., 2000)
and more recently have been proposed as a new bacterial taxon
(Bianciotto et al., 2003). Endocellular bacteria were found in
the glomeromycotean associates in nearly all the liverwort taxa
examined by electron microscopy. The spheroidal shape,
absence of a bounding fungal membrane, and relatively thick
cell walls of the gram-positive type distinguish these bacteria
from those in the Gigasporaceae. Bacterial endophytes similar
to those in liverwort-associated glomeromycotean fungi have
been reported in Glomus fistulosum in an artificial association
with the hornwort Anthoceros punctatus (Schu¨ ßler, 2000); in
Geosiphon pyriforme, a glomeromycotean fungus associated
with a cyanobacterium (Schu¨ ßler et al., 1994); and in putative
glomeromycotean fungi associated with wild gametophytes of
several basal taxa including the hornwort Phaeoceros laevis
(Ligrone, 1988), the lycopod Lycopodium clavatum (Schmid
and Oberwinkler, 1993), and the eusporangiate ferns Bo-
trychium (Schmid and Oberwinkler, 1994) and Tmesipteris
(Duckett and Ligrone, 2005).
Concluding remarks—This study confirms the widespread
occurrence of glomeromycotean associations in basal liverwort
lineages and suggests that these associations involve cellular
and molecular interactions comparable in complexity to those
in AMs (Paszkowski, 2006). The results support the hypothesis
that the two associations are homologous in terms of biological
interactions (Wang and Qiu, 2006) but do not provide an
unequivocal answer as to which of them is ancestral. The basal
position of the liverworts in embryophyte phylogeny and the
widespread occurrence of GAs in basal liverwort clades are
consistent with the view that coevolution of glomeromycotean
fungi with liverworts preceded the appearance of AMs in
tracheophytes (Kottke and Nebel, 2005; Wang and Qiu, 2006).
This view gains support also from the presence of closely
similar associations in the gametophytes of lycopods (Duckett
and Ligrone, 1992; Schmid and Oberwinkler, 1993), basal
ferns (Schmid and Oberwinkler, 1994, 1995; Duckett and
Ligrone, 2005), and hornworts (Ligrone, 1988; Schu¨ ßler,
2000), the last now proposed as the sister group to the
tracheophytes (Fig. 11) based on recent molecular and
immunocytochemical data (Dombrovska and Qiu, 2004;
Carafa et al., 2005; Groth-Malonek et al., 2005; Qiu et al.,
2006). However, the finding that the fungal endophytes in a
number of liverwort taxa, taxa widely separated both
phylogenetically and geographically, are all related to the
Glomus Group A is what one might expect under the
hypothesis of host shifting from tracheophytes to liverworts
(Selosse, 2005). The two models are not mutually exclusive: in
a tracheophyte-dominated world, advanced glomeromycotean
fungi from tracheophytes should be expected to replace more
primitive endophytes in adapted potential hosts. Further field
and molecular research and resynthesis experiments might help
solve this interesting evolutionary issue.
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APPENDIX. Voucher information for liverwort taxa used in this study. Voucher specimens are deposited in the following herbaria: BM ¼British Museum;
DGL ¼private herbarium, D. G. Long, Royal Botanical Garden, Edinburgh; JGD ¼private herbarium, J. G. Duckett at Queen Mary, University of
London.
Asterella bachmanii JGD Jan 1995 South Africa.
A. australis JGD Oct Nov Dec 1999 JGD Sept 2001 New Zealand.
A. tenera JGD Oct Nov Dec 1999 JGD Sept 2001 New Zealand.
A. muscicola JGD Jan 1995 Lesotho.
A. wilmsii JGD Jan 1995 Lesotho, JGD Jan 1992 South Africa.
Aitchisoniella himalayensis BM July 1933 India.
Allisonia cockaynii JGD Sept Oct 1999 Jan 2000 Sept 2001 New Zealand.
Aneura lobata subsp. australis JGD Sept Oct 1999 Aug Sept 2001 New
Zealand.
A. maxima JGD Apr 2007 USA.
A. novaeguineensis JGD Jan 2000.
A. pinguis JGD Aug 1983 Sept 2005 11 Nov 2006 8 Dec 2006 UK.
A. pseudopinguis JGD June 1989 Jan 1995 Lesotho.
Apometzgeria pubescens JGD 11 Nov 2006 4 Apr 2004 UK.
Athalamia hyalina JGD June 2004 Italy, JGD 3 Aug 2005 USA.
A. pinguis DGL 30889 India.
Austrofossombronia australis JGD Sept 1999 New Zealand.
Blasia pusilla JGD Oct 1980 JGD Aug 1990 JGD 11 Nov 2006 JGD 2 Feb
2007 UK, JGD Aug 1995 USA.
Bucegia romanica BM 10 Sept 1940 BM 19 Nov 1909 Rumania.
Cyathodium cavernarum JGD Aug 1998 Uganda.
C. foetidissimum JGD June 2003 Italy.
Conocephalum conicum JGD Aug 1972 France, JGD 5 Nov 1996 24 Feb
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS

2006 Italy, JGD 8 Apr 1965 JGD 24 July 1967 JGD 1 Apr 1973 JGD
21 Jan 2002 JGD 2 Apr 2004 UK, JGD 28 Mar 2007 JGD 3 Apr 2007
USA.
C. salebrosum JGD 11 Nov 2006 JGD 8 Dec 2006 UK, JGD 28 Mar 2007
JGD 3 Apr 2007 USA.
Corsinia coriandra JGD 3 Nov 1996 JGD 24 Feb 2006 Italy.
Cronisia fimbriata BM 8903 Brazil.
Cryptomitrium oreoides JGD Jan 1994 JGD Jan 1995 Lesotho.
Cryptothallus mirabilis JGD 12 Feb 1967 JGD 11 Mar 1967 JGD Apr
1996 JGD Sept 1998 UK.
Dumortiera hirsuta JGD 12 Sept 2006 Chile, JGD July 1973 France, JGD
18 May 2005 Venezuela, JGD Aug 1966 UK.
Exomotheca holstii JGD Jan 1995 Lesotho.
E. pustulosa JGD Jan 1994 Lesotho.
F. caespitiformis JGD 3 Nov 1996 24 Feb 2006 Italy.
F. echinata JGD 24 Feb 2006 Italy.
F. maritima JGD Sept 1972 UK.
F. pusilla JGD 11Apr 1969 JGD 4 Apr 1974 Sept 2006 UK.
F. wondraczeckii JGD 16 Aug 1968 JGD Nov 1972 JGD 2 Nov 1975 UK.
Fossombronia angulosa JGD 2 Jan 2007 France, JGD 2 Feb 2006 Italy,
JGD 22 Mar 1966 JGD 2 Apr 1970 JGD Apr 1978 UK.
Geothallus tuberosus JGD Aug 1995 USA.
Greeneothallus gemmiparus JGD 19 Jan 2005 Chile.
Haplomitrium blumei JGD June 1995 JGD Feb 2000 Malaysia.
H. chilensis JGD 16 17 &19 Jan 2005 Chile.
H. gibbsiae JGD Oct Nov Dec 1999 JGD Jan Feb 2000 JGD Sept 2001
Oct 2001 New Zealand, JGD Aug 1998 Uganda.
H. hookeri JGD 24 & 26 Aug 1968 JGD 25 Sept 1982 JGD 10 Aug 1996
JGD 11 Nov 2006 UK.
H. intermedium JGD Aug 1981 Australia.
H. ovalifolium JGD Jan 2000 JGD Sept 2001 New Zealand.
Hymenophyton flabellatum JGD Sept Nov 1999 JGD Jan 2000 JGD Sept
2001 New Zealand.
Jensenia connivens JGD 18 May 2005 Venezuela.
J. wallichii JGD 18 May 2005 Venezuela.
Lunularia cruciata JGD 2 Jan 2007 France, JGD 23 Feb 2006 JGD 2 Nov
1996 Italy, JGD Jan 1982 JGD Sept 1968 JGD 10 Jan 2007 UK.
Moerckia blyttii JGD Aug 2003 Switzerland, JGD 19 July 1967 JGD 9
Aug 1968 JGD 26 Aug 1968 JGD Sept 1984 UK.
M. hibernica JGD 8 Dec 2006 JGD Feb 2007 UK.
Mannia angrogyna JGD 25 Feb 2006 Italy.
M. fragrans DGL 27059 China, JGD 28 Oct 2005 Germany.
Marchantia berteroana JGD 12 Aug 2006 Chile, JGD 18 May 2005
Venezuela.
M. foliacea JGD 9 Jan 2005 Chile, JGD Jan 2000 JGD Sept 2001 New
Zealand.
M. pappeana JGD Jan 1991 JGD Jan 1995 Lesotho.
Marchantia polymorpha subsp. polymorpha JGD 24 Aug 1966 JGD 15
Apr 1967 JGD 23 Aug 1969 JGD 19 June 2007 UK.
M. polymorpha subsp. ruderalis JGD Sept 1994 JGD 20 Sept 1999 JGD
10 June 2007 UK.
M. polymorpha subsp. montivagans JGD11 Nov 2006 JGD 8 Dec 2006
UK.
Metzgeria conjugata JGD Aug 1964 JGD 11 Nov 2006 JGD 2 Feb 2007
UK.
M. decipiens JGD Jan 2005 JGD Aug 2006 JGD Sept 2006 Chile.
M. fruticulosa JGD 2 Mar 2005 JGD 12 Feb 2006 UK.
M. furcata JGD Aug 1964 JGD 12 Feb 2006 JGD 26 July 2006 JGD I0
Jan 2007 UK.
M. temperata JGD Sept 2006 JGD May 2007 UK.
Monocarpus sphaerocarpus JGD Aug 1981 BM June 1971 Australia.
Monoclea forsteri JGD Oct Nov Dec 1999 JGD Jan Feb 2000 JGD Sept
Oct 2001 New Zealand.
M. gottschei JGD 12 16 Sept 2006 Chile, JGD June 1998 Mexico, 16 May
2005 Venezuela.
Monosolenium tenerum JGD 28 Oct 2005 Germany (from aquarium), JGD
Nov 2006 Japan.
Neohodgsonia mirabilis JGD Jan 2000 Sept 2001 New Zealand.
Noteroclada confluens JGD 9 Jan 2005 JGD 19 Jan 2005 12 Sept 2006
Chile, JGD 18 May 2005 Venezuela.
Oxymitra cristata JGD Jan 1992 Lesotho.
Oxymitra incrassata JGD 26 Feb 2006 Italy.
Pallavicinia connivens JGD Nov 1999 JGD Sept 2001 New Zealand.
P. indica JGD Aug 1981 Malaysia.
P. lyellii JGD Nov 2006 UK, JGD Apr 2007 USA.
P. tenuinervis JGD Nov 1999 JGD Sept 2001 New Zealand.
P. xiphoides JGD Dec 1999 Sept 2001 New Zealand.
Pellia endiviifolia JGD 22 Feb 2006 Italy, JGD Apr 1983 JGD 4 Apr 2004
JGD Dec 2005 JGD 8 Dec 2006 UK.
P. epiphylla JGD 4 Apr 2004 JGD Sept 2006 JGD 8 Dec 2006 2 Feb 2007
UK, JGD 20 Mar 5 2007 JGD 3 Apr 2007 USA.
P. neesiana JGD Nov 1972 JGD 6 Sept 1974 JGD 2 Feb 2007 UK.
Peltolepis grandis BM July 1882 Norway, BM 2 Aug 1876 Russia
(Siberia), BM Aug 1906 Switzerland.
Petalophyllum ralfsii JGD Feb 2003 Italy, JGD Mar 1968 JGD Aug 1979
JGD 8 Dec 2006 UK.
Phyllothallia nivicola JGD 17 Jan 2005 Chile, JGD Jan 2000 New
Zealand.
Plagiochasma exigua JGD Jan 1992 JGD Jan 1995 South Africa, JGD Jan
1993 JGD Jan 1995 Lesotho.
P. rupestre JGD Jan 1992 JGD Jan 1994 JGD Jan 1995 South Africa, JGD
Jan 1989 JGD Jan 1996 Lesotho.
Pleurozia purpurea JGD 22 Aug 1966 JGD July 1996 JGD 2 Feb 2007
UK.
P. gigantea JGD June 1995 Malaysia.
Podomitrium phyllanthus JGD Oct 1999 New Zealand.
Preissia quadrata JGD 28 Feb 2006 Italy, JGD 6 Apr 1973 JGD Aug
1979 JGD 11 Nov 2006 JGD 8 Dec 2006 UK.
Reboulia hemispherica JGD 15 Jan 2005 JGD 8 Sept 2006 Chile, JGD
May 2003 JGD 23 Feb 2006 Italy, JGD 27 Aug 1964 JGD 3 Apr 2004
JGD 8 Dec 2006 UK.
Riccardia chamedryfolia JGD 7 Apr 1967 JGD 9 Apr 1968 JGD Oct 2005
JGD 2 Feb 2007 UK.
R. cochleata JGD Oct 1999 New Zealand.
R. eriocaula JGD Oct 1999 JGD Sept 2001 New Zealand.
R. incurvata JGD 12 Apr 1968 JGD 14 Aug 1968 JGD 8 Dec 2006 JGD 2
Feb 2007 UK.
Riccardia intercellula JGD Sept 2001 New Zealand.
R. latifrons JGD 26 Aug 1966 JGD 6 Apr 1967 JGD 2 Feb 2007 UK.
R. multifida JGD 6 Apr 1967 JGD 8 Aug 1968 JGD 2 Feb 2007 UK.
R. pennata JGD Sept 2001 New Zealand.
Riccia albolimbata JGD 24 Nov 2005 Botswana.
R. beyrichiana JGD 8 May 1971 UK.
R. canaliculata JGD 10 Nov 1972 JGD 1 Aug 1978 UK.
R. cavernosa JGD June 1989 JGD Jan 1994 Lesotho, 22 Oct 1967 JGD 12
Oct 1969 JGD 16 Sept 1970 UK.
R. crozalsii JGD 22 Feb 2006 Italy, JGD 19 Mar 1968 JGD June 2004
UK.
R. crystallina JGD Jan 1994 Lesotho, JGD 6 May 1968 JGD June 1989
UK.
R. fluitans JGD 1 Dec 1968 JGD 12 Oct 1969 JGD 7 Dec 1969 JGD Dec
2006 UK.
R. glauca JGD Apr 1972 JGD Sept 1994 JGD Apr 2003 JGD Nov 2005
UK.
R. huebeneriana JGD 1 Dec 1968 UK.
R. montana JGD Jan 1995 Lesotho.
R. nigrella JGD 24 Feb 2006 Italy, JGD June 1989 JGD Jan 1995 Lesotho,
JGD Sept 2001 New Zealand, JGD Apr 1967 JGD 19 Mar 1968 UK.
R. okahandjana JGD 24 Nov 2005 Botswana.
R. sorocarpa JGD 18 Mar 1968 JGD 11 Nov 2006 UK.
R. stricta JGD 23 Nov 2005 Botswana, JGD June 1989 JGD Jan 1995
Lesotho.
R. subbifurca JGD June 1968 JGD Sept 2004 JGD Nov 2006 UK.
Ricciocarpus natans JGD 16 May 1966 JGD 3 Sept 1967 UK.
Riella americana JGD Aug 1995 USA.
R. helicophylla JGD Aug 1970 Greece.
AMERICAN JOURNAL OF BOTANY [Vol. 94

Symphyogyna brasiliensis JGD Jan 1995 South Africa, JGD 18 May 2005
Venezuela.
S. brogniartii JGD 18 May 2005 Venezuela.
S. hymenophyton JGD Oct Nov 1999 JGD Aug Sept 2001 New Zealand.
S. subsimplex JGD Oct 1999 JGD Sept 2001 New Zealand.
S. undulata JGD Oct 1999 JGD Sept 2001 New Zealand.
Sauteria alpina BM 30 June 1870 June 1880 Switzerland.
Sphaerocarpos michelii JGD Nov 1996 Italy, JGD 7 Apr 6 May 1968 UK.
S. texanus JGD 6 May 1968 UK.
Stephensoniella brevipedunculata BM Nov 1934 DGL 30890 India.
Targionia hypophylla JGD 4 Apr 1967 JGD 28 Dec 2006 France, JGD 5
Nov 1996 JGD 24 Feb 2006 Italy, JGD Oct 1999 JGD Feb 2000 JGD
Sept 2001 New Zealand, JGD 5 May 1968 JGD 23 Mar 1969 UK.
Treubia lacunosa JGD Sept 2001 New Zealand.
T. lacunosoides JGD Sept Oct 1999 JGD Jan Feb 2000 JGD Sept Oct
2001 New Zealand.
T. pygmaea JGD Oct Nov 1999 JGD Jan 2000 JGD Sept Oct Nov 2001
New Zealand.
Verdoornia succulenta JGD Jan 2000 JGD Sept 2001 New Zealand.
Wiesnerella denudata BM Apr 1951 Japan, BM 27 July 1953 Java, DGL
30673 Nepal, BM 11 Apr 1899 Sikkim.
Xenothallus vulcanicolus JGD Oct 2001 New Zealand.
November 2007] LIGRONE ET AL.—GLOMEROMYCOTEAN ASSOCIATIONS IN LIVERWORTS