R L A C E L V B P B J G. D · 2008-03-06 · only occur in tracheophytes but also in the...

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. DUCKETT4

2Dipartimento di Scienze ambientali, Seconda Universita di Napoli, via A. Vivaldi 43, I-81100 Caserta, Italy;3Dipartimento 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; and4School 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 isconsidered a major factor involved in the colonization ofterrestrial habitats by phototrophic organisms (Selosse and LeTacon, 1998). It is assumed that the common ancestor to theGlomeromycota, Ascomycota, and Basidiomycota originatedafter the appearance of land plants (Berbee and Taylor, 2007)and that the association with glomeromycotean fungi, to formthe so-called arbuscular mycorrhizas (AMs), is a plesiomorphy(primitive character) in the tracheophytes. Already present inSiluro-Devonian fossils of protracheophytes and still occurring

in the majority of present-day tracheophytes (Selosse and LeTacon, 1998; Wang and Qiu, 2006), the AMs have beenreplaced by associations with basidio- or ascomycetes inseveral derived lineages of higher plants (Wang and Qiu, 2006;Berbee and Taylor, 2007). Endophytic fungal associations notonly occur in tracheophytes but also in the gametophytes ofliverworts and hornworts, while they appear to be absent inmosses (Read et al., 2000; Renzaglia et al., 2007).

The fungal associations in members of the Marchantiopsida(complex thalloid liverworts) and Metzgeriidae (simple thalloidliverworts) are cytologically similar to AMs (Strullu et al.,1981; Pocock and Duckett, 1984; Ligrone and Lopes, 1989;Ligrone and Duckett, 1994). Similar associations have alsobeen described in Haplomitrium and Treubia (Carafa et al.,2003; Duckett et al., 2006a), two taxa recently placed in a cladethat 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 fungalsymbiont in Marchantia foliacea has been identified asbelonging to the glomeromycotean genus Glomus, group A(Russell and Bulman, 2005). An assemblage of simple thalloidliverworts and the leafy liverworts (Jungermanniidae) form adiversity 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 asrevealed by recent molecular work (Davis, 2004; Forrest andCrandall-Stotler, 2004, 2005; Heinrichs et al., 2005), it hasbeen suggested that the association with glomeromycoteanfungi is a plesiomorphy in the liverworts (Nebel et al., 2004;Kottke and Nebel, 2005). Moreover, considering that theliverworts are almost unanimously recognized as the earliest-

1 Manuscript received 24 February 2007; revision accepted 16 August2007.

This work was funded by grants from the Seconda Universita di Napoliand Regione Campania, Italy (LR 5, 2003). The research in Torino wasfunded 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 atthe IMAGE Center (Southern Illinois University), and the staff at theCISME (University of Naples ‘‘Federico I,’’ Italy) for laboratory andelectron 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 ZealandDepartment 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 thisstudy. J.G.D. was supported by an overseas travel grant from the RoyalSociety (UK) in New Zealand and by a DEFRA Darwin Initiative grant inChile. R.L. was supported by a grant from CNR (Italy) in New Zealand.

5 Author for correspondence (e-mail: [email protected])

American Journal of Botany 94(11): 1756–1777. 2007.

divergent clade in the phyletic tree of land plants (Nickrent etal., 2000; Dombrovska and Qiu, 2004; Groth-Malonek et al.,2005; Qiu et al., 2006) and that the Glomeromycota are basal tothe other mycorrhiza-forming fungi (Schußler et al., 2001;James et al., 2006), it has been suggested that glomeromyco-tean associations (GAs) in liverworts predated the arbuscularmycorrhizas in vascular plants (Nebel et al., 2004; Kottke andNebel, 2005; Duckett et al., 2006a; Wang and Qiu, 2006). Analternative scenario, i.e., secondary host shift of glomeromy-cotean symbionts from tracheophytes to liverworts, has beenconsidered by Selosse (2005), mainly on the basis of Russelland Bulman’s (2005) identification of the fungal endophyte ofMarchantia paleacea as a member of the Glomus Group A,i.e., a derived group in the phyletic tree of Glomeromycota(Schußler et al., 2001).

In spite of the growing interest in fungus–liverwortassociations in recent years, current information on theircytology and physiology is remarkably sparse. In particular, asconcerns putative GAs, the information available for most ofthe taxa reported by Nebel et al. (2004) is from lightmicroscopy and generally does not go beyond the notion ofthe presence/absence of fungal endophytes tentatively referredto as glomero, basidio- or ascomycetes. Owing to the smallnumber of taxa investigated in detail to date, it is impossible toreach any general conclusions about the cytology of putativeGAs in liverworts.

The general aim of this long-standing investigation was toprovide an exhaustive survey of the biology of GAs inliverworts and specifically to (1) identify the fungal endophytesthrough molecular analysis in selected liverwort taxa; (2)determine the taxonomic and geographical distribution of GAsin liverworts through a morphological (light and electronmicroscopy) analysis of taxa collected worldwide; (3) inves-tigate the level of cellular compatibility between liverworts andfungi through a detailed immunocytochemical analysis of theircontact surfaces; and (4) confirm Koch’s postulates through invitro synthesis of GAs from axenic liverwort cultures andspores of known glomeromycotean fungi. The data presentedare discussed in the context of the origins of GAs in liverwortsand their evolutionary relationships with AMs.

MATERIALS AND METHODS

The liverwort species examined, their taxonomic position, fungal status, andgeographical origin are listed in Table 1. Liverwort taxonomy follows Crandall-Stotler and Stotler (2000) and Heinrichs et al. (2005). With the exception of fewexceedingly rare species, the diagnosis for fungal status was based on the studyof samples from at least two separate collection sites and from freshly-collectedspecimens. At least 20 plants were examined for each sample. For voucherinformation of the taxa examined in this study, see the Appendix.

Molecular analysis—Molecular analysis of fungal endophytes was carriedout for the following liverwort species: Haplomitrium chilensis, Conocephalumconicum, 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 colonizedparts were isolated with a razor blade under a dissecting microscope. Thesamples, each about 50–100 mg, were surface-sterilized with cloramine T (3%)and streptomycin (0.3%) followed by two rounds of sonication. A mininum oftwo 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 universaleukaryotic 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 asnegative controls.

The PCR reaction was performed in a total volume of 25 lL containing 2 lLof template solution, 0.2 mM of each dNTP, 10 pmols of each primer, 1 U ofREDTaq DNA polymerase (Sigma, St. Louis, Missouri, USA) and 13 REDTaqReaction buffer (SIGMA). Amplification was performed in a GeneAmp PCRsystem 9700 (PerkinElmer, Waltham, Massachussets, USA) programmed asfollows: 1 3 3 min at 958C; 35 3 1 min at 958C, 1 min at 588C, 2 min at 758C;1 3 7 min at 728C. Electrophoretical analysis of the PCR products revealed asingle band of 550 bp. This fragment was purified from gel using the QIAquickpurification kit (QIAGEN), cloned into a pGEM-T Easy Vector (Promega,Madison, Wisconsin, USA), and then transformed into Escherichia coli JM109High Efficiency Competent Cells (Promega).

Thirty putatively positive transformant clones (white colonies) from eachliverwort sample were selected manually, and the DNA extracted from eachclone was amplified using the PCR mix and program detailed previously. ForRFLP (restriction fragment length polymorphism) analysis, aliquots of 4 lL ofeach PCR amplicon were mixed with 16 lL of digestion mix containing 2.0 lLbuffer 103, 0.2 lL bovine serum albumin, 13.3 lL H2O, and 0.5 lL of therestriction enzyme Hinf I or Hsp92II (Promega) for 3 h at 378C. Fragmentpatterns were analyzed on agarose gel containing 0.84 % agarose (Sigma) and1.5% high-resolution agarose (Sigma). One to four PCR amplicons weresequenced for each restriction pattern and species, using the vector-specificprimers T7 and SP6, at the DNA Sequences Naples Facilities. The sequenceshave been deposited in GenBank under the accession numbers reported inTable 3.

Forward and reverse sequences were analyzed using the program BioLign4.0.6 (http://en.bio-soft.net/dna/BioLign.html). DNA sequences were comparedto GenBank database using the BLAST algorithm (Altschul et al., 1997) foridentification. Data bank sequences with high homology to our sequences wereincluded in the data set, using the profile alignment function CLUSTAL W(Thompson et al., 1994) for multiple alignment. The nearest relatives of eachsequence were inferred with the neighbor-joining algorithm (Saitou and Nei,1987) and the Kimura two-parameter model (Kimura, 1980), using the PHYLIPpackage (Felsenstein, 1989). The confidence of branching was assessed using1000-bootstrap resampling (Felsenstein, 1985).

Light and electron microscopy—Both fresh and fixed samples wereexamined by light microscopy. The visibility of fungal hyphae in rhizoids andhand-cut sections of the thallus was improved by staining with 0.05% trypanblue in lactophenol (Ligrone and Lopes, 1989) or 0.05% aniline blue in lacticacid. Autofluorescence of liverwort cell walls was observed on fresh hand-cutsections using an excitation filter at 365 nm and a barrier filter with atransmission cutoff at 397 nm.

Colonized areas of the thallus of fungus-containing specimens were cut intosmall pieces under a dissecting microscope and fixed with a mixture of 3%glutaraldehyde, 1% freshly prepared formaldehyde, and 0.75% tannic acid in0.04 M piperazine-N,N0–bis(2-ethanesulfonic acid) (PIPES) buffer, pH 7.0, for2 h at room temperature under gentle vacuum. The samples were then rinsed in0.08 M PIPES buffer and twice in 0.08 M Na-cacodylate buffer, and postfixedin 1% OsO4 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 propyleneoxide at 48C, the samples were slowly infiltrated with Spurr’s resin(Polysciences, Warrington, Pennsylvania, USA) at 48C, transferred topolypropylene dishes, and cured at 688C for 24 h. For light microscopy, 0.5-lm-thick sections of resin-embedded samples were cut with a diamondhistoknife, stained with 0.5% toluidine blue O in 1% Na-tetraborate, andphotographed with a Zeiss Axioskop (Zeiss, Jena, Germany) light microscopeequipped 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% methanolfor 15 min and in Reynold’s lead citrate for 10 min, and observed with a Jeol1200 EX2 (Jeol, Tokyo, Japan) electron microscope.

For scanning electron microscopy (SEM), the samples were cut with a razorblade and taken through a 1 : 1 ethanol:acetone series to remove the cytoplasm,osmicated for 48 h in aqueous 2% OsO4, and stored in 70% ethanol. Thesamples were then dehydrated in anhydrous ethanol and critical point driedusing CO2 as the transfusion fluid, mounted on stubs, and sputter-coated with390 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) Neesa G UK (5)H. intermedium Berrieac G Australia (1)H. ovalifolium R.M. Schusta G New Zealand (2)H. chilensis R.M. Schust.ac G Chile (3)

TreubialesTreubia lacunosa (Colenso) Prosk.a G New Zealand (1)T. lacunosoides Pfeiffer, Frey & Stechaac 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 Bellardic — Italy (1), UK (1)S. texanus Austinc — UK (1)Geothallus tuberosus Campb.bc — USA (1)Riella americana M.Howe & Underwoodc — USA (1)Riella helicophylla (Boryet Mont.) Mont.c — Greece (1)

MonoclealesMonoclea forsteri Hook.a G New Zealand (7)M. gottschei Lindb.a G Chile (2), Mexico (1), Venezuela (1)

MarchantialesAytoniaceae

Asterella bachmanii (Steph.) S.W.Arnellac G South Africa (1)A. muscicola (Steph.) S.W.Arnellc G Lesotho (1)A. wilmsii (Steph.) S.W.Arnellac 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 Peroldc — Lesotho (2)Mannia angrogyna (L.) A. Evansa — 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.) Raddia G Italy (2), UK (3), Chile (2)

WiesnerellaceaeWiesnerella denudata Schiffn.b — (2) Japan (1), Java (1), Nepal (1), Sikkim (1)

ConocephalaceaeConocephalum conicum (L.) Dumort.a G France (1), Italy (2), UK (6), USA (2)C. salebrosum Szweykowski, Buczkowska & Odrzykoskic G UK (2), USA (3)

LunulariaceaeLunularia cruciata (L.) Dumort. ex Lindb.a G France (1), Italy (2), UK (4)

MarchantiaceaeBucegia romanica Raddibc — Rumania (2)Dumortiera hirsuta (Sw.) Neesa 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. & Boisseliera G UK (2)Neohodgsonia mirabilis (H. Perss.) H. Perss.a G (3) New Zealand (2)Preissia quadrata (Scop.) Neesa G Italy (1), UK (4)

MonosoleniaceaeMonosolenium tenerum Griffithc — Germany (from aquarium) (1), Japan (1)Peltolepis grandis Lindb.b — Norway (1), Russia (Siberia) (1), Switzerland (1)

CleveaceaeAthalamia hyalina (Sommerf.) S. Hatt.c G Italy (1), USA (1)A. pinguis W. Falc.bc G India (1)Sauteria alpina (Nees) Neesb — Switzerland (2)

ExormothecaceaeAitchisoniella 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 Kunzec — Uganda (1)

AMERICAN JOURNAL OF BOTANY [Vol. 94

TABLE 1. Continued.


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)

MonocarpaceaeMonocarpus sphaerocarpus Carrc — Australia (1)

TargionaceaeTargionia hypophylla L.a G France (2), Italy (2), New Zealand (3), UK (2)

OxymitraceaeOxymitra incrassata (Broth.) Sergio & Sim-Sima — Italy (1)O. cristata Garside ex Peroldc — Lesotho (1)

RicciaceaeRiccia subgenus RicciellaR. 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.) Peroldc — Lesotho (2), Botswana (1)Riccia subgenus RicciaR. albolimbata S.W.Arnellc — Botswana (1)R. beyrichiana Hampe ex Lehm. — UK (1)R. crozalsii Levierc — Italy (1), UK (2)R. glauca L. — UK (4)R. montana Peroldc — Lesotho (1)R. nigrella DC.c — Italy (1), Lesotho (2), New Zealand (1), UK (2)R. okahandjana S.W.Arnellc — 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 MacVicarac 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 & Gottschea G Italy (1), UK (3)

AllisoniaceaeAllisonia cockaynii (Steph.) R.M. Schust.ac G New Zealand (4)

PelliaceaeNoteroclada confluens Tayl. ex Hook. & Wilsona G Chile (3), Venezuela (1)Pellia endiviifolia (Dicks.) Dum.a G Italy (1), UK (4)P. epiphylla ( L.) Cordaa G UK (5), USA (2)P. neesiana (Gottsche) Limpr. G UK (4)

PallaviciniaceaeGreeneothallus gemmiparus Hasselac G Chile (1)Jensenia connivens (Colenso) Grolleac G Venezuela (1)J. wallichii Colensoac G Venezuela (1)Moerckia hibernica (Hook.) Gottschea — (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.) Graya — (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. & Neesc G New Zealand (4)S. subsimplex Mitt.c G New Zealand (2)S. undulata Colensoc G New Zealand (2)Xenothallus vulcanicolus R.M. Schust.ac G New Zealand (1)

HymenophytaceaeHymenophyton flabellatum (Labill.) Dum.a G (1) New Zealand (4)


Immunocytochemistry—Epitopes associated with cell wall polysaccha-

rides and proteins were localized immunocytochemically in Marchantiapolymorpha 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.


MakinoaceaeVerdoornia succulenta R.M. Schust.ac B New Zealand (2)

AneuraceaeAneura lobata subsp. australis R.M. Schust.ac B New Zealand (4)A. maxima Schiffn. (Steph.)a B (3) USA (1)A. novaeguineensis Hewsonac B New Zealand (1)A. pinguis (L.) Dum.a B UK (4)A. pseudopinguis (Herzog) Pocsac B Lesotho (2)Cryptothallus mirabilis Malmba B UK (4)Riccardia intercellula E.A.Brownc B New Zealand (1)R. pennata E.A.Brownc B New Zealand (1)R. chamedryfolia (With.) Grolle — (3) UK (4)R. cochleata (Hook.f. & Taylor) Kuntzec — 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)

MetzgeriaceaeApometzgeria 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.) Duma — UK (4)M. fruticulosa (Dicks.) A. Evans — UK (2)

PleuroziaceaePleurozia 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 oforigin 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, thefungal 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-bpamplicons 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) andHsp92II (S1-S4), and GenBank sequence codes (in parentheses) offungal small-subunit rDNA 550-bp amplicons from fungus-associatedliverworts.

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)


night temperature regime. After 3 mo in culture, plates containing thalli about10 mm long were inoculated with glomeromycotean spores that had beenpreviously surface-sterilized with 3% chloramine T and 0.03% streptomycin for5 min. Four different glomeromycotean species were tested: Gigaspora roseaNicolson & 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 Glomusclarum Nicolson & Schenck (BEG142), kindly supplied by Dr. VivienneGianinazzi-Pearson (INRA, Dijon, France). The cultures were examined atintervals with a dissection microscope. Putative associations, identified fromfungal colonization of rhizoids and of the internal tissue of the thalli, wereprocessed for light and electron microscopy as described.

RESULTS

Molecular identification of fungal endophytes—PCRamplification of DNA from fungus-colonized liverwort tissuewith the NS31 and AM1 primers produced a DNA fragment ofabout 550 bp. RFLP analysis of this fragment with therestriction enzymes Hinf I and Hsp92II produced six and fourdifferent RFLP types, respectively (Table 2). One to threedifferent restriction patterns were obtained from each liverwortspecies, and for each pattern one to four amplicons weresequenced (Table 3). When the sequences were aligned with adata set from GenBank, they all clustered within theGlomeromycota with high bootstrap support, producing a treetopology coherent with those from Schußler et al. (2001) (Fig.1). The sequence from Monoclea was closely related withAcaulospora, while the remaining sequences were related withthe Glomus Group A, in part clustering within this group and inpart forming a sister clade to it (Fig. 1).

The taxonomic distribution of glomeromycotean associa-tions in liverworts—The cytology of the GAs for the fungalendophytes that were identified by molecular analysis (see nextsection) 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 pathogenicsymptoms in host plants, intracellular colonization by aseptatehyphae, fungal colonization restricted to specific tissue areas inthe gametophyte (see next section) and absent from thesporophyte, fungal entry via the rhizoids (except in the

Haplomitriopsida, the development of intracellular arbuscule-like structures, the development of fungal vesicles, andendobacteria occurring in fungal hyphae. Septate fungi wereidentified by electron microscopy as basidiomycetes orascomycetes from the presence of dolipores or simple septa,respectively.

While in the majority of GA-forming taxa the fungalassociation was consistently present regardless of the collectingsite or season, in a few species the degree of colonization washighly variable from plant to plant even within the samesample. For example, populations of Monoclea forsteri, M.gottschei, Conocephalum conicum, Lunularia cruciata, Du-mortiera hirsuta, and Noteroclada confluens growing either invery wet or epilithic habitats were more variable than werepopulations growing on soil. In Marchantia polymorpha, thefungus was present in the subspecies montivagans, a perennialtaxon growing in natural habitats, but was absent from the twopioneer subspecies, polymorpha and ruderalis, that colonizeephemeral and usually nutrient-rich habitats. Also lackingendophytes were the linear branches with very few rhizoids inPellia spp. from wet habitats and the furcate caducous rhizoid-free branches of P. endiviifolia that proliferate in the autumnand early winter (Paton, 1999). For each liverwort taxonexamined, GAs were reported as present when consistentlyfound in at least a part of the specimens examined, providedthat the morphological criteria detailed previously weresatisfied. Our reports refer to the potential ability, or apparentinability (the latter amenable to confutation by examination offurther samples), of certain taxa to establish this type ofsymbiosis, with no assumption of ecological relevance.Moreover, no attempt was made in this study to quantitativelyevaluate the occurrence of the fungi.

Based on the guidelines described, GAs were found to bewidespread in a large liverwort assemblage encompassing theHaplomitriopsida, Marchantiopsida, and part of the Metzger-iidae within the Jungermanniopsida (Table 1). Within theMarchantiopsida, fungal endophytes were consistently absentin 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, AustraliaCCRC-M1 Xyloglucan, rhamnogalacturonan-I/terminal (1!2)-a-linked

fucosyl-containing side groupPuhlman et al., 1994/M. Hahn, Complex Carbohydrate Research Center,

University of Georgia, USACCRC-M2 Rhamnogalacturonan-I/unknown Puhlman et al., 1994/M. Hahn, Complex Carbohydrate Research Center,

University of Georgia, USACCRC-M7 Arabinogalactan-proteins, rhamnogalacturonan-I/

arabinosilated (1!6)-b-galactanPuhlman et al., 1994; Steffan et al., 1995/M. Hahn, Complex Carbohydrate

Research Center, University of Georgia, USALM1 Hydroxyproline-rich glycoproteins/undefined Smallwood et al., 1996/J.P. Knox, Centre for Plant Sciences,

University of Leeds, UKLM2 Arabinogalactan-proteins/undefined Smallwood et al., 1995/J.P. Knox, Centre for Plant Sciences,

University of Leeds, UKLM5 Galactan, rhamnogalacturonan-I/tetra (1!4)-b-galactan Jones et al., 1997/J.P. Knox, Centre for Plant Sciences, University of Leeds, UKLM6 Arabinan, rhamnogalacturonan-I/penta(1!5)-a-arabinan Willats et al., 1998/J.P. Knox, Centre for Plant Sciences, University of Leeds, UKLM7 Homogalacturonan/partially methyl esterified Willats et al., 2001/J.P. Knox, Centre for Plant Sciences, University of Leeds, UKJIM5 Homogalacturonan/low- or unmethyl-esterified Willats et al., 2000/J.P. Knox, Centre for Plant Sciences, University of Leeds, UKJIM7 Homogalacturonan/partially methyl-esterified Willats et al., 2000/J.P. Knox, Centre for Plant Sciences, University of Leeds, UKJIM11 Hydroxyproline-rich glycoproteins/undefined Smallwood et al., 1994/J.P. Knox, Centre for Plant Sciences,

University of Leeds, UK


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


iidae clade I (Davis, 2004), GAs were common, with theexception of a few isolated species in the Pallaviciniaceae,while the remaining taxa traditionally included in theMetzgeriidae and grouped in the Metzgeriidae clade II byDavis (2004) were either fungal free (Pleuroziaceae andMetzgeriaceae) or associated with basidiomycetes (Aneuraceaeand Verdoornia). In no case has a putative GA been detected inthe 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 theMarchantiopsida and Metzgeriidae, including the speciesinvestigated by molecular techniques, glomeromycotean colo-nizations were typically localized in the rhizoids and theinternal 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 placentalarea associated with the foot, were never colonized. The oil-body idioblasts in marchantialean liverworts remained fungalfree 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) Marchantiapolymorpha subsp. montivagans, and (D) Symphyogyna brasiliensis, showing colonized areas (F) of the internal parenchyma. Scale bars: A–D, 100 lm.


the midrib in certain marchantialean taxa such as Conocepha-lum and Marchantia (Fig. 2C). In many members of thePallaviciniaceae, GAs were restricted to subterranean stolonslacking a laminar margin (Fig. 2D). Fungi were also rare orabsent in the lipid-laden regions of the perennating tubers ofPetalophyllum ralfsiii and Fossombronia maritima.

Unlike the Haplomitriopsida (Carafa et al., 2003; Duckett etal., 2006a), direct fungal penetration through epidermal cellswas never observed in the Marchantiopsida or Metzgeriidae,indicating that here the rhizoids are the only access to thefungus. The fungus penetrated the rhizoids at any point andformed large intracellular hyphae running in both directions(Fig. 3A, B). Of the two types of rhizoids present in manymarchantialean liverworts, i.e., living smooth rhizoids andtuberculate rhizoids that undergo cytoplasmic lysis at maturity,

only the former were found to be primarily colonized byglomeromycotean fungi. From rhizoids, the hyphae entered theparenchyma cells above the lower epidermis of the thallus (Fig.3C, D). The colonization was entirely intracellular and closelyresembled Paris-type arbuscular mycorrhizas (Smith andSmith, 1997), with large colonizing hyphae spreading fromcell to cell and intercalary formation of arbuscule-likestructures from shorter lateral branches, or trunk hyphae, withdeterminate growth (Fig. 3E, F). In taxa with large thalli, suchas Conocephalum conicum or Marchantia polymorpha, thecolonizing hyphae often grew longitudinally along the midribof the thallus, probably extending the colonization to asignificant distance from the entry point (Fig. 4A). In mostother taxa, however, the hyphae had no particular orientation inthe 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-colonizedrhizoid of Conocephalum conicum; the arrow points to the penetration site of a fungal hypha. (B) Rhizoid of Marchantia polymorpha subsp. montivaganscontaining fungal hyphae and vesicles (arrows). (C) Rhizoid base with fungal hypae (arrow) in Monoclea forsteri. (D) Fungal hyphae (arrow) passing fromthe rhizoid base (R) to adjacent parenchyma cells in Preissia quadrata. (E) Colonizing hypha crossing host cell walls (arrows) and (F) arbuscule in thethallus parenchyma of Fossombronia echinata. Scale bars: A–D, 20 lm; E, F, 10 lm.


association was found in the genus Marchantia. Here thefungus colonized a region overarching the midrib hyalinestrand and consisting of a lower area containing coils andvesicles and an upper area containing arbuscules (Fig. 4B–D).Cell wall autofluorescence, a normal feature of fungus-freeparenchyma cells in the thallus of Conocephalum and othermarchantialean liverworts, was no longer visible in colonizedcells (Fig. 5A). Autofluorescence disappeared first from thecell 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 locallysis, while the host plasmalemma invaginated to form aperifungal membrane that surrounded the intracellular fungus

and separated it from the host cytoplasm. An interfacial matrixof fibrillar material was deposited in the space between thehyphae and perifungal membrane, with a thickness decreasingfrom 0.5–1.0 lm at entry/exit points, where it formed aconspicuous collar around the fungus (Fig. 5D), to 50 nm orless in fine arbuscular hyphae (Fig. 6E). The colonizing hyphaewere 3–6 lm in diameter, rarely less, and had relatively thickwalls (100–200 nm), sometimes with a layered structure (Fig.6A); the fungal cell walls retained the same thickness orbecame slightly thinner in larger trunk hyphae, but they thinnedto about 30 nm or less in terminal arbuscular hyphae (Fig. 6E).

The colonizing hyphae contained numerous vacuoles withelectron-transparent contents and irregular shapes; scattered inthe 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 showinglarge colonizing hyphae (arrows) growing along the longitudinal axis of the thallus. (B) Colonized area in the thallus midrib of Marchantia polymorphasubsp. 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) lowerand (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.


brane-bound spheroidal bodies of electron-opaque material(Figs. 5D and 6A, B). The trunk hyphae of the arbusculesappeared similar to colonizing hyphae, but often they could bedistinguished because their cytoplasm was filled with minutevacuoles (Fig. 5E). With very few exceptions (e.g., the fungalendophytes in Haplomitrium gibbsiae and Pellia epiphylla),endocellular bacteria were present in both colonizing and trunkhyphae. These appeared as cocci about 0.3–0.5 lm in diameter,sometimes of more irregular shape, with an electron-opaquecell wall of the gram-positive type (i.e., relatively thick andlacking an outer membrane) and no bounding fungalmembrane. Division of bacterial endophytes by a central

constriction was observed frequently (Fig. 6C). The terminalarbuscular hyphae typically were less than 1 lm in diameterand contained no nuclei nor endobacteria (Fig. 6E). As reportedfor Treubia (Duckett et al., 2006a), the fungal endophyte inPetalophyllum did not form typical arbuscules but only coiledhyphae of relatively uniform diameter (Fig. 6D). Thecolonizing hyphae in Petalophyllum did not exceed 3 lm indiameter, and the thinner intracellular hyphae were rarely lessthan 1 lm.

With fungal colonization, the host cells underwent pro-nounced morphological changes that were remarkably uniformin 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-colonizedarea (F ). The red fluorescence is from chloroplasts in the adjacent chlorenchyma. (B) Bright-field micrographs of colonizing hypha spreading in the thallusparenchyma of C. conicum; the arrows point to host cell walls crossed by the fungus. (C) Fluorescence micrographs of the same area showing thatautofluorescence first disappears in cell walls crossed by the fungus (arrows). (D, E) Transmission electron micrograph of fungus-colonized thallusparenchyma cells in Marchantia polymorpha subsp. montivagans. (D) Detail of colonizing hypha crossing a host cell wall; the arrows point to the collarsof interfacial material; fungal nucleus (N ); perifungal membrane continuous with host plasmalemma (arrowheads). (E) Detail of an arbuscule-containingcell, 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.


Fig. 6. Transmission electron micrographs of glomeromycotean associations in liverworts. (A) Transverse section of a colonizing hypha in Pelliaepiphylla; the arrow points to the thick multilayered wall. (B) Detail of colonizing hypha in P. epiphylla showing membrane-bound, electron-opaquebodies (arrows). (C) Bacterial endophyte with central constriction in a trunk hypha in Marchantia paleacea; note the gram-positive type bacterial wall andthe 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 includingmitochondria (M ), microbodies (Mb), and plastids (P). Note the absence of starch in plastids. A dictyosome (G ) and several profiles of endoplasmicreticulum 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.


Fig. 7. Transmission electron micrographs of glomeromycotean associations in liverworts. (A) Oil bodies (OB) in fungus-free and (B) colonizedparenchyma cell in Petalophyllum ralfsii, showing differences in the appearance of the matrix and lipid components; note the starch-filled plastids (P) inthe fungus-free cell. (C) Intravacuolar clump of collapsed hyphae in Hymenophyton flabellatum. (D) Degenerated hyphae (F ) in P. ralfsii; the arrow pointsto 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 muchthicker and the cytoplasm is packed with lipid (L). Scale bars: A, C, 1 lm; B, D, 0.5 lm; E, F, 2 lm.


cytoplasm and organelles, replacement of the large centralvacuole typical of fungus-free cells with numerous smallervacuoles separated by cytoplasmic strands, migration of thenucleus from a peripheral to an internal position, anddisappearance or strong reduction of starch in plastids. Thenucleus and plastids often became pleomorphic. The fungusestablished intimate spatial relationships with the nucleus,plastids, mitochondria, and other host organelles (Figs. 5E and6E). In the Metzgeriopsida (in which all somatic cells typicallycontain oil bodies) the density of the matrix and the abundanceof lipid droplets in the oil bodies markedly declined incolonized cells (Fig. 7A, B).

At a more advanced stage of colonization, the arbusculesdegenerated, forming one to several clumps of collapsedhyphae. The larger hyphae usually survived the arbuscules andcould occasionally give rise to a second colonization cycle,inferred from the presence of a healthy arbuscular system alongwith clumps of collapsed hyphae in the same cells. When allthe intracellular fungus was dead, the host cells resumed theirprecolonization cytological organization, the only sign of pastcolonization being intravacuolar clumps of fungal wall remains(Fig. 7C). A different pattern of fungal degeneration wasobserved in Petalophyllum. Here the hyphae underwent cellwall dissolution and cytoplasmic lysis, producing masses ofamorphous material in which no fungal walls were discernible(Fig. 7D). Common in degenerated hyphae in Petalophyllumwere ghosts of crystals, probably calcium oxalate, thatdissolved during fixation (Fig. 7D).

Vesicles developed by terminal swelling of colonizinghyphae or of lateral branches. Initial vesicle development wascharacterized by nuclear and organelle proliferation (Fig. 7E);at later stages the vesicles accumulated abundant lipid reservesand their cell walls thickened conspicuously (Fig. 7F).

Immunocytochemistry—The immunocytochemical tests inC. 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 fungalpenetration nor in the interfacial material covering theintracellular hyphae. The same antibody also labeled the wallof hyphae external to the thallus (Fig. 8B) but not ofintracellular hyphae; in the latter some labeling was observedonly within the vacuoles (Fig. 8C). JIM5 and JIM7, twoantibodies against homogalacturonan, and JIM11, whichrecognizes an epitope associated with hydroxyprolyne-richproteins, labeled the liverwort cell walls throughout (except thecell corners) as well as the interfacial material associated withthe intracellular fungus (Fig. 8D–F).

Perhaps the most interesting results were those obtained withCCRC-M1, a monoclonal antibody that recognizes fucosylatedside groups associated with xyloglucan and rhamnogalactur-onan I (Puhlman et al., 1994). This antibody produced verylittle labeling of the cell walls in fungus-free cells, includingmeristematic cells. In contrast, the same antibody stronglylabeled the interfacial material associated with the intracellularfungus (Fig. 9A, B). Moreover, starting from the penetrationsite, colonized cells deposited a new wall layer that wascontinuous with the interfacial material and was also heavilylabeled by CCRC-M1 (Fig. 9C).

In vitro synthesis of glomeromycotean associations—Thefour fungal isolates tested were all able to colonize the roots ofthe higher plant Trifolium repens L., producing typical AMs. Incontrast, successful colonization of the host liverwort (C.conicum) was obtained only with spores of Glomus mosseaeand only in about 10% of the plants inoculated. In the othercases, the fungal spores either failed to germinate (Glomusclarum) or produced germlings that stopped growing and died(Gigaspora rosea and G. margarita, and some G. mosseaespores).

The colonized plants were maintained in culture for severalmonths 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 inexternal hyphae

þþþ Plasmodesmatal collars þþ Cell walls inexternal hyphae

� Rest of cell walls � Cell walls ininternal hyphae

� Rest of cell walls � Cell walls ininternal hyphae

þ Vacuoles þ VacuolesCCRC-M1 6 Cell walls in fungus-free cells 6 Cell walls in fungus-free cells

þþ Cell walls in colonized cellsand interfacial material

� þþþ Cell walls in colonized cellsand 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, þ weak, 6 very weak and uneven, – absent. Positive reports with no furtherinformation indicate nonspecific labeling of all host cell walls and of the interfacial material.


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 liverwortspecies selected for molecular analysis, three were from Europe(Conocephalum, Fossombronia, and Pellia), one was fromNew Zealand (Monoclea), and one was from South America(Haplomitrium). Molecular analysis demonstrates that thesespecies all contain fungal endophytes that cluster with theGlomeromycota 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 Conocephalumconicum. (A) Labeling of the host cell wall around the plasmodesmata (arrows), indicating the presence of callose. (B) Labeling of the fungal wall inexternal hyphae (arrows). (C) Detail of a colonizing hypha at the penetration point; no labeling is visible in the fungal wall (FW) or in the interfacialmaterial (IM ); some labeling is visible in fungal vacuoles (FV ); host cell wall (HW ). (D–F) Details of colonizing hyphae (F ) and host cell wall atpenetration points in Marchantia polymorpha subsp. montivagans, showing labeling with (D) JIM7, (E) JIM5, and (F) JIM11; these antibodies labeledboth the host walls (HW ) and interfacial material (IM ). Scale bars: A–F, 0.3 lm.


Acaulospora. Both glomeromycotean lineages form arbuscularmycorrizas in tracheophytes (Smith and Read, 1997; Petersonet al., 2004). The results are consistent with a former study thatdemonstrated the presence of glomeromycotean endophytesrelated to Glomus Group A in populations of Marchantiafoliacea in New Zealand (Russell and Bulman, 2005). In linewith molecular analysis, our resynthesis experiments showedthat G. mosseae, a glomeromycotean fungus that nests withinthe Glomus Group A (Fig. 1), was able to colonize axenic thalliof C. conicum and to establish an endophytic associationclosely similar to that observed in the wild. A similar result wasobtained in a cross-colonization experiment with the simplethalloid liverwort Pellia epiphylla and an unidentifiedglomeromycotean fungus associated with the higher plantPlantago lanceolata (Read et al., 2000). The low frequency ofcolonization 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 lowcompatibility and/or an inhibitory effect of growth conditionson the liverwort ability to elicit fungal development. In linewith the first possibility is the repeated occurrence of the samefungal phylotypes in populations of M. paleacea from differentsites (Russell and Bulman, 2005). Although too few taxa havebeen studied to support any general conclusion, the datasuggest a degree of specificity between liverworts and GlomusGroup A that contrasts with the large spectrum of glomero-mycotean associates in tracheophytes (Peterson et al., 2004).

The taxonomic distribution and origins of GAs inliverworts—The application of diagnostic criteria inferred

Fig. 9. Immunogold labeling with CCRC-M1 monoclonal antibody inglomeromycotean associations in liverworts. (A) Colonizing hypha (F )crossing a host cell wall (HW ) in Marchantia polymorpha subsp.montivagans; the antibody labeled the interfacial material covering thefungus (arrows). (B) Detail of (A), showing heavy labeling of interfacialmaterial at the fungal entry point (arrow). (C) Detail of a cell wall at theinterface between a fungus-colonized (CC ) and a fungus-free (UC ) hostcell; a heavily labeled cell wall layer (bracket) is visible on the sidetowards the colonized cell while no labeling is visible on the other side ofthe cell wall. Scale bars: A–C, 0.3 lm.

Fig. 10. Resynthesis of a glomeromycotean association from spores ofGlomus mosseae and axenic thalli of Conocephalum conicum. (A) Handsection of a thallus stained with aniline blue, showing fungus-colonizedrhizoids (arrows) and parenchyma cells (IP). (B) Transmission electronmicrographs of colonized parenchyma cell, showing profiles of colonizinghypha (CH ), trunk hyphae (TH ), and arbuscular hyphae (AH ). Scale bars:A, 40 lm; B, 3 lm.


from the cytological analysis of the liverwort species withfungal endophytes that we identified by molecular techniqueshas provided more solid support for the morphologicalidentification of GAs in other taxa. With information on 67species with a previously unknown fungal status and thereexamination of 64 species already included in the list byNemec et al. (2004), our survey confirms GAs as a generalfeature of a large liverwort assemblage encompassing theHaplomitriopsida, most of the Marchantiopsida, and part of theMetzgeriidae (the simple thalloid clade I according to Davis,2004). With reference to the topology of the phyletic tree ofliverworts 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 taxonomicdistribution strongly suggests that the symbiotic associationwith glomeromycotean endophytes is a plesiomorphy inliverworts. Accordingly, the consistent absence of GAs incertain taxa, both basal (Blasiales and Sphaerocarpales) andderived (Ricciaceae, the simple thalloid clade II and the wholeclade of leafy liverworts) should be interpreted as the result ofmultiple independent losses. However, the apparent liverworttendency to associate predominantly with fungi related to theGlomus Group A is consistent with host shifting of symbiontsfrom tracheophytes to liverworts (Selosse, 2005). The latterhypothesis might explain in terms of multiple acquisitions, atleast in part, the scattered distribution of GAs in liverworts.

Discrepancies between the present study and the survey byNebel et al. (2004) relative to certain taxa, in particularCorsinia coriandrina and Hymenophyton flabellatum (Table1), may reflect intraspecific ecological variability. Further

investigation is needed to ascertain whether the absence offungal endophytes in several isolated species within mycorrh-ized families, such as Cryptomitrium oreoides in theAytoniaceae or several species of Pallavicinia in thePallaviciniaceae, are further instances of multiple evolutionaryloss/acquisition or of ecological variability as noted inConocephalum, Lunularia, Pellia, Noteroclada, Dumortiera,and Monoclea. As in vascular plants, many of the liverwortsthat lack GAs grow in very wet habitats. Paradoxically,however, absence is equally common in liverwort taxa growingin places subjected to intense seasonal desiccation. The absenceof GAs from the two Marchantia polymorpha subspeciesgrowing in nutrient-rich habitats (polymorpha and ruderalis) isnot unexpected and suggests that shifting from the mycorrhizalto nonmycorrhizal status in liverworts is relatively easy inevolutionary terms.

Our survey confirms the absence of GAs in the Pleuro-ziaceae and Metzgeriaceae, and we report the presence ofbasidiomycetous endophytes not only in the Aneuraceae butalso in Verdoornia, a taxon traditionally placed in the distantlyrelated family Makinoaceae (Crandall-Stotler and Stotler,2000). In molecular phylogenies these four groups form asingle clade (simple thalloid II, Fig. 11) with a sisterrelationship to the leafy liverworts (Davis, 2004; Forrest andCrandall-Stotler, 2004; Heinrichs et al., 2005). More detailedanalysis is now needed to ascertain possible affinities of thebasidiomycete associations in Verdoornia and in the Aneur-aceae (Kottke et al., 2003) and thereby to gain insight into theevolution of these associations following the postulated loss ofGAs in the common ancestor to the simple thalloid II/leafyliverwort 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 theHaplomitriopsida have several unique features including thecolonization of epidermal cells in Haplomitrium, the coloni-zation of intercellular spaces in Treubia, and the developmentof thin-walled hyphal swellings in both genera (Carafa et al.,2003; Duckett et al., 2006a). Because molecular analysis hasshown that the fungal endophyte of H. chilensis clusters withthe endophytes from marchantialean and metzgerialaleanliverworts, the distinctive morphology of GAs in theHaplomitriopsida appears to depend on control by the hostrather than the fungus.

The results of immunocytochemical analysis of GAs inConocephalum and Marchantia indicate a level of functionalinteraction between the symbionts comparable to that in AMs.No callose deposition was observed in colonized cells at thepoints of fungal entry nor at the host/fungus interface. Callosedeposition has been implicated in numerous studies as aresistance response to attack by pathogens (Rodriguez-Galvezand Mendgen, 1995; Enkerli et al., 1997), while higher plantsproduce little or no callose in reacting to AM fungi (Balestriniet al., 1994; Gianinazzi-Pearson et al., 1996). Also significantis the observation that the antibody against (1!3)-b-glucanlabels the cells walls of external hyphae but not those ofintracellular hyphae, suggesting that the association with thehost liverwort inhibits the synthesis of this polysaccharide inthe fungus. A decrease in cell wall labeling by antibodiesagainst (1!3)-b-glucan in AM fungi has been interpreted as asign of structural simplification of the fungal wall accompa-

Fig. 11. Phyletic tree of the liverworts and their position relative to therest of the embryophytes. The asterisks indicate the clades that formendophytic associations with glomeromycetes. For further details aboutliverwort 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).


nying the development of the intraradical phase (Lemoine etal., 1995).

The cell walls in the thallus parenchyma of Marchantia andConocephalum were strongly labeled by antibodies againsthomogalacturonans with different degrees of methyl esterifi-cation (JIM5 and JIM7) and by an antibody that recognizes anepitope associated with hydroxyprolyne-rich proteins (JIM11).Both groups of compounds are widespread components of cellwalls in plants but are not known in fungal walls. Therefore,the presence of the same epitopes in the interfacial matrixensheathing the intracellular mycobiont indicates that, as inAMs (Balestrini et al., 1996; Harrison, 1997; Balestrini andBonfante, 2005), this material is of host origin and that the hostcells colonized by the fungus maintain the ability to synthesizeand secrete cell wall material. The results obtained with CCRC-M1 demonstrate that the fungal colonization elicits thesynthesis of cell wall polysaccharide(s) that are scarcelypresent in fungus-free thallus parenchyma cells. The suppres-sion of autofluorescence in colonized cells also indicateschanges in cell wall composition consequent to fungalcolonization. No change in the expression of the CCRC-M1epitope like that observed in this study has been reported inother glomeromycotean associations. Immunogold labeling ofAMs in higher plants with CCRC-M1 and CCRC-M7 showedthat, although the tissue distribution of the epitopes of thesetwo antibodies varied according to the plant species, theinterfacial matrix invariably had the same labeling pattern asthat found in host cell walls before fungal colonization(Balestrini et al., 1996). In contrast, fungal colonization incucumber AMs elicited the expression of two differentexpansin proteins, one localized in the host cell walls and theother in the interfacial matrix (Balestrini et al., 2005).

Endocellular bacteria are common in glomeromycoteanfungi forming AMs in higher plants. Originally reported as‘‘bacterium-like organelles,’’ the glomeromycotean endobac-teria were first studied by Macdonald et al. (1982), whodescribed three different types, either free in fungal cytoplasmor 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 inthe glomeromycotean associates in nearly all the liverwort taxaexamined by electron microscopy. The spheroidal shape,absence of a bounding fungal membrane, and relatively thickcell walls of the gram-positive type distinguish these bacteriafrom those in the Gigasporaceae. Bacterial endophytes similarto those in liverwort-associated glomeromycotean fungi havebeen reported in Glomus fistulosum in an artificial associationwith the hornwort Anthoceros punctatus (Schußler, 2000); inGeosiphon pyriforme, a glomeromycotean fungus associatedwith a cyanobacterium (Schußler et al., 1994); and in putativeglomeromycotean fungi associated with wild gametophytes ofseveral basal taxa including the hornwort Phaeoceros laevis(Ligrone, 1988), the lycopod Lycopodium clavatum (Schmidand Oberwinkler, 1993), and the eusporangiate ferns Bo-trychium (Schmid and Oberwinkler, 1994) and Tmesipteris(Duckett and Ligrone, 2005).

Concluding remarks—This study confirms the widespreadoccurrence of glomeromycotean associations in basal liverwortlineages and suggests that these associations involve cellular

and molecular interactions comparable in complexity to thosein AMs (Paszkowski, 2006). The results support the hypothesisthat the two associations are homologous in terms of biologicalinteractions (Wang and Qiu, 2006) but do not provide anunequivocal answer as to which of them is ancestral. The basalposition of the liverworts in embryophyte phylogeny and thewidespread occurrence of GAs in basal liverwort clades areconsistent with the view that coevolution of glomeromycoteanfungi with liverworts preceded the appearance of AMs intracheophytes (Kottke and Nebel, 2005; Wang and Qiu, 2006).This view gains support also from the presence of closelysimilar associations in the gametophytes of lycopods (Duckettand Ligrone, 1992; Schmid and Oberwinkler, 1993), basalferns (Schmid and Oberwinkler, 1994, 1995; Duckett andLigrone, 2005), and hornworts (Ligrone, 1988; Schußler,2000), the last now proposed as the sister group to thetracheophytes (Fig. 11) based on recent molecular andimmunocytochemical 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 anumber of liverwort taxa, taxa widely separated bothphylogenetically and geographically, are all related to theGlomus Group A is what one might expect under thehypothesis of host shifting from tracheophytes to liverworts(Selosse, 2005). The two models are not mutually exclusive: ina tracheophyte-dominated world, advanced glomeromycoteanfungi from tracheophytes should be expected to replace moreprimitive endophytes in adapted potential hosts. Further fieldand molecular research and resynthesis experiments might helpsolve 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: BM¼British Museum;DGL¼ private herbarium, D. G. Long, Royal Botanical Garden, Edinburgh; JGD¼ private herbarium, J. G. Duckett at Queen Mary, University ofLondon.

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 NewZealand.

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


2006 Italy, JGD 8 Apr 1965 JGD 24 July 1967 JGD 1 Apr 1973 JGD21 Jan 2002 JGD 2 Apr 2004 UK, JGD 28 Mar 2007 JGD 3 Apr 2007USA.

C. salebrosum JGD 11 Nov 2006 JGD 8 Dec 2006 UK, JGD 28 Mar 2007JGD 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 2006Chile, 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 2004JGD 8 Dec 2006 UK.

Riccardia chamedryfolia JGD 7 Apr 1967 JGD 9 Apr 1968 JGD Oct 2005JGD 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.


Symphyogyna brasiliensis JGD Jan 1995 South Africa, JGD 18 May 2005Venezuela.

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 JGDSept 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.


R L A C E L V B P B J G. D · 2008-03-06 · only occur in tracheophytes but also in the...

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