| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
What's this? |
Anatomy and Morphology |
Dipartimento di Scienze ambientali, Seconda Università di Napoli, via A. Vivaldi 43, I-81100 Caserta, Italy; Dipartimento di Biologia vegetale, Università 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; School of Biological and Chemical Sciences, Queen Mary University of London, Mile End Road, London E1 4NS, UK
Received for publication February 24, 2007. Accepted for publication August 16, 2007.
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.
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 basidiomycetes 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-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 (Schüßler et al., 2001
; James et al., 2006
), it has been suggested that glomeromycotean 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 glomeromycotean 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 (Schüßler 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) investigate 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.
|
) 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 µL containing 2 µL 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 1x REDTaq Reaction buffer (SIGMA). Amplification was performed in a GeneAmp PCR system 9700 (PerkinElmer, Waltham, Massachussets, USA) programmed as follows: 1 x 3 min at 95°C; 35 x 1 min at 95°C, 1 min at 58°C, 2 min at 75°C; 1 x 7 min at 72°C. 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 µL of each PCR amplicon were mixed with 16 µL of digestion mix containing 2.0 µL buffer 10x, 0.2 µL bovine serum albumin, 13.3 µL H2O, and 0.5 µL of the restriction enzyme HinfI or Hsp92II (Promega) for 3 h at 37°C. 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.
|
|
) 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,N'–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% OsO4 in 0.08 M Na-cacodylate buffer, pH 6.7, overnight at 4°C. Following dehydration in a step gradient of ethanol and one step in propylene oxide at 4°C, the samples were slowly infiltrated with Spurr's resin (Polysciences, Warrington, Pennsylvania, USA) at 4°C, transferred to polypropylene dishes, and cured at 68°C for 24 h. For light microscopy, 0.5-µm-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% OsO4, and stored in 70% ethanol. The samples were then dehydrated in anhydrous ethanol and critical point dried using CO2 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.
Immunocytochemistry
Epitopes associated with cell wall polysaccharides 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, Pennsylvania, USA), and cured at 60°C 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.
|
). 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 12/10°C day/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 HinfI 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 Schüßler 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).
|
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, Dumortiera 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 Metzgeriidae 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, Monocarpaceae, Oxymitraceae, and Ricciaceae. Within the Metzgeriidae 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 colonizations 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 the midrib in certain marchantialean taxa such as Conocephalum 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.
|
; 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 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).
|
|
|
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 µm 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 µm 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 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 µm in diameter, and the thinner intracellular hyphae were rarely less than 1 µm. With fungal colonization, the host cells underwent pronounced morphological changes that were remarkably uniform in all taxa investigated. These included proliferation of the 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).
|
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)-ß–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).
|
|
). 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).
|
The colonized plants were maintained in culture for several months with no adverse symptoms, although fungal colonization did not appreciably enhance their growth relative to the 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 parenchyma 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).
|
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 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 glomeromycotean 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 glomeromycotean associates in tracheophytes (Peterson et al., 2004
).
The taxonomic distribution and origins of GAs in liverworts
The application of diagnostic criteria inferred 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.
|
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 mycorrhized 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 Pleuroziaceae 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 Aneuraceae (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 Marchantiopsida and Metzgeriidae are remarkably uniform in development and morphology. In contrast, GAs in the Haplomitriopsida have several unique features including the colonization of epidermal cells in Haplomitrium, the colonization 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 (13)-ß-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 (13)-ß-glucan in AM fungi has been interpreted as a sign of structural simplification of the fungal wall accompanying 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 esterification (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 suppression 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 endobacteria 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 Gigasporaceae have been identified as gram-negative ß-proteobacteria 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 (Schüßler, 2000
); in Geosiphon pyriforme, a glomeromycotean fungus associated with a cyanobacterium (Schüßler 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 Botrychium (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
; Schüßler, 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.
APPENDIX
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 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.
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.
FOOTNOTES
1 This work was funded by grants from the Seconda Università 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{at}unina2.it
)
LITERATURE CITED
Altschul S. F. Madden T. L. Schäffer A. A. Zhang J. Zhang Z. Miller W. Lipman D. J.. 1997. Gapped BLAST and PSI-BLAST, a new generation of protein database search programs. Nucleic Acids Research 25: 3389-3402..
Balestrini R. Bonfante P.. 2005. The interface compartment in arbuscular mycorrhizae: a special type of plant cell wall?. Plant Biosystems 139: 8-15..
Balestrini R. Cosgrove D. J. Bonfante P.. 2005. Differential location of 
–expansin proteins during the accommodation of root cells to an arbuscular mycorrhizal fungus. Planta 220: 889-899..[CrossRef][Web of Science][Medline]
Balestrini R. Hahn M. G. Faccio A. Mendgen K. Bonfante P.. 1996. Differential localization of carbohydrate epitopes in plant cell walls in the presence and absence of arbuscular mycorrhizal fungi. Plant Physiology 111: 203-213..[Abstract]
Balestrini R. Romera C. Puigdomenech P. Bonfante P.. 1994. Localization of a cell-wall hydroxyproline-rich glycoprotein, cellulose and ß-1,3-glucans in apical and differentiated regions of maize mycorrhizal roots. Planta 195: 201-209..[Web of Science]
Berbee M. L. Taylor J. W.. 2007. Rhynie chert: a window into a lost world of complex plant–fungus interactions. New Phytologist 174: 475-479..[CrossRef][Web of Science][Medline]
Bianciotto V. Lumini E. Bonfante P. Vandamme P.. 2003. ‘Candidatus Glomeribacter gigasporarum’ gen. nov., sp. nov., an endosymbiont of arbuscular mycorrhizal fungi. International Journal of Systematic and Evolutionary Microbiology 53: 121-124..
Bianciotto V. Lumini E. Lanfranco L. Minerdi D. Bonfante P. Perotto S.. 2000. Detection and identification of bacterial endosymbionts in arbuscular mycorrhizal fungi belonging to the family Gigasporaceae. Applied and Environmental Microbiology 66: 4503-4509..
Carafa A. Duckett J. G. Knox J. P. Ligrone R.. 2005. Distribution of cell-wall xylans in bryophytes and tracheophytes: new insights into basal relationships of land plants. New Phytologist 168: 231-240..[CrossRef][Web of Science][Medline]
Carafa A. Duckett J. G. Ligrone R.. 2003. Subterranean gametophytic axes in the primitive liverwort Haplomitrium harbour a unique type of endophytic association with aseptate fungi. New Phytologist 160: 185-197..[CrossRef][Web of Science]
Crandall-Stotler B. Stotler R.. 2000. Morphology and classification of the Marchantiophyta. In A. J. Shaw, B. Goffinet [eds.], Bryophyte biology, 21-70. Cambridge University Press, Cambridge, UK..
Davis E. C.. 2004. Molecular phylogeny of leafy liverworts according to analysis of twelve genes. In B. Goffinet, V. Hollowell, R. Magill [eds.], Molecular systematics of bryophytes, 61-68. Missouri Botanical Garden Press, St. Louis, Missouri, USA..
Dombrovska O. Qiu Y. L.. 2004. Distribution of introns in the mitochondrial gene nad1 in land plants: phylogenetic and molecular evolutionary implications. Molecular Phylogenetics and Evolution 32: 246-263..[CrossRef][Web of Science][Medline]
Duckett J. G. Carafa A. Ligrone R.. 2006a. A highly differentiated glomeromycotean association with the mucilage-secreting antipodean liverwort Treubia; clues to the origins of mycorrhizas. American Journal of Botany 93: 797-813..
Duckett J. G. Ligrone R.. 1992. A light and electron microscope study of the fungal endophytes in the sporophyte and gametophyte of Lycopodium cernuum with observations of the gametophyte–sporophyte junction. Canadian Journal of Botany 70: 58-72..
Duckett J. G. Ligrone R.. 2005. A comparative cytological analysis of fungal endophytes in the sporophyte rhizomes and vascularized gametophytes of Tmesipteris and Psilotum. Canadian Journal of Botany 83: 1443-1456..[CrossRef]
Duckett J. G. Russell J. Ligrone R.. 2006b. Basidiomycetous endophytes in jungermannialean (leafy) liverworts have novel cytology and species-specific host ranges: a cytological and experimental study. Canadian Journal of Botany 84: 1075-1093..[CrossRef]
Enkerli K. Hahn M. G. Mims C. W.. 1997. Immunogold localization of callose and other plant cell wall components in soybean roots infected with the oomycete Phytophthora sojae. Canadian Journal of Botany 75: 1509-1517..[CrossRef]
Felsenstein J.. 1985. Confidence limits on phylogenies: an approach using the bootstrap. Evolution 39: 783-791..[CrossRef][Web of Science]
Felsenstein J.. 1989. PHYLIP: Phylogeny Inference Package, version 3.2. Cladistics 5: 164-166..
Forrest L. A. Crandall-Stotler B. J.. 2004. A phylogeny of the simple thalloid liverworts (Jungermanniopsida, subclass Metzgeriidae) as inferred from five chloroplast genes. In B. Goffinet, V. C. Hollowell, R. E. Magill [eds.], Molecular systematics of bryophytes, 119-140. Missouri Botanical Garden Press, St. Louis, Missouri, USA..
Forrest L. A. Crandall-Stotler B. J.. 2005. Progress towards a robust phylogeny for the liverworts, with particular focus on the simple thalloids. Journal of the Hattori Botanical Laboratory 97: 127-159..
Forrest L. L. Davis E. C. Long D. G. Crandall-Stotler B. J. Clark A. Hollingsworth M. L.. 2006. Unraveling the evolutionary history of the liverworts (Marchantiophyta): multiple taxa, genomes and analyses. Bryologist 109: 303-334..[CrossRef]
Gianinazzi-Pearson V. Dumas-Gaudot E. Gollotte A. Tahiri-Alaoui A. Gianinazzi S.. 1996. Cellular and molecular defense-related root responses to invasion by arbuscular mycorrhizal fungi. New Phytologist 133: 45-57..[CrossRef][Web of Science]
Groth-Malonek M. Pruchner D. Grewe F. Knoop V.. 2005. Ancestors of trans-splicing mitochondrial introns support serial sister group relationships of hornworts and mosses with vascular plants. Molecular Biology and Evolution 22: 117-125..
Harrison M. J.. 1997. The arbuscular mycorrhizal symbiosis: an underground association. Trends in Plant Science 2: 54-60..[CrossRef][Web of Science]
Heinrichs J. Gradstein S. R. Wilson R. Schneider H.. 2005. Towards a natural classification of liverworts (Marchantiophyta) based on the chloroplast gene rbcL. Cryptogamie, Bryologie 26: 131-150..[Web of Science]
Helgason T. Daniell T. J. Husband R. Fitter A. H. Young J. P. W.. 1998. Ploughing up the wood-wide web?. Nature 394: 431..[CrossRef][Medline]
James T. Y. et al (71 authors).. 2006. Reconstructing the early phylogeny of fungi using a six-gene phylogeny. Nature 443: 818-822..[CrossRef][Medline]
Jones L. Seymour G. B. Knox J. P.. 1997. Localization of pectic galactan in tomato cell walls using a monoclonal antibody specific to (14)-ß-d-Galactan. Plant Physiology 113: 1405-1412..[Abstract]
Kimura M.. 1980. A simple method for estimating evolutionary rate of base substitutions through comparative studies of nucleotide sequences. Journal of Molecular Evolution 16: 111-120..[CrossRef][Web of Science][Medline]
Kottke I. Beiter A. Weiß B. M. Haug I. Oberwinkler F. Nebel M.. 2003. Heterobasidiomycetes form symbiotic associations with hepatics. Jungermanniales have sebacinoid mycobionts while Aneura pinguis (Metzgeriales) is associated with Tulasnella species. Mycological Research 107: 957-968..[CrossRef][Web of Science][Medline]
Kottke I. Nebel M.. 2005. The evolution of mycorrhiza-like associations in liverworts: an update. New Phytologist 167: 330-334..[CrossRef][Web of Science][Medline]
Lemoine M. C. Gollotte A. Gianinazzi-Pearson V.. 1995. Localization of ß-(13) glucan in walls of the endomycorrhizal fungi Glomus mosseae (Nicol. & Gerd.) Gerd. & Trappe and Acaulospora laevis Gerd. & Trappe during colonization of host roots. New Phytologist 129: 97-105..[CrossRef][Web of Science]
Ligrone R.. 1988. Ultrastructure of a fungal endophyte in Phaeoceros laevis (L.) Prosk. (Anthocerotophyta). Botanical Gazette 149: 92-100..
Ligrone R. Duckett J. G.. 1994. Thallus differentiation in the marchantialean liverwort Asterella wilmsii (Steph.) with particular reference to longitudinal arrays of endoplasmic microtubules in the inner cells. Annals of Botany 73: 577-586..
Ligrone R. Lopes C.. 1989. Cytology and development of a mycorrhiza-like infection in the gametophyte of Conocephalum conicum (L.) Dum. (Marchantiales, Hepatophyta). New Phytologist 111: 423-433..[CrossRef][Web of Science]
Ligrone R. Vaughn K. C. Renzaglia K. S. Knox J. P. Duckett J. G.. 2002. Diversity in the distribution of polysaccharide and glycoprotein epitopes in the cell walls of bryophytes: new evidence for multiple evolution of water-conducting cells. New Phytologist 156: 491-508..[CrossRef][Web of Science]
Macdonald R. M. Chandler M. R. Mosse B.. 1982. The occurrence of bacterium-like organelles in vesicular-arbuscular fungi. New Phytologist 90: 659-663..[CrossRef][Web of Science]
Meikle P. J. Bonig I. Hoogenraad N. J. Clarke A. E. Stone B. A.. 1991. The location of (13)-ß-glucans in the walls of pollen tubes of Nicotiana alata using a (13)-ß-glucan-specific monoclonal antibody. Planta 185: 1-8..[Web of Science]
Murashige T. Skoog F.. 1962. A revised medium for rapid growth and bio-assays with tobacco tissue cultures. Physiologia Plantarum 15: 473-497..[CrossRef]
Nebel M. Kreier H. P. Preussing M. Weiß M. A. Kottke I.. 2004. Symbiotic fungal associations with liverworts are the possible ancestors of mycorrhizae. In R. Agerer, H. Piepenbring, P. Blanz [eds.], Frontiers in basidiomycete mycology, 339-360. HIW-Verlag, Ecking, Germany..
Nickrent D. L. Parkinson C. L. Palmer J. D. Duff R. J.. 2000. Multigene phylogeny of land plants with special reference to bryophytes and the earliest land plants. Molecular Biology and Evolution 17: 1885-1895..
Paszkowski U.. 2006. A journey through signaling in arbuscular mycorrhizal symbioses 2006. New Phytologist 172: 35-46..[CrossRef][Web of Science][Medline]
Paton J. A.. 1999. The liverwort flora of the British Isles. Harley Books, Colchester, UK..
Peterson R. L. Massicotte H. B. Melville L. H.. 2004. Mycorrhyzas: anatomy and cell biology. NRC Research Press, Ottawa, Quebec, Canada..
Pocock K. Duckett J. G.. 1984. A comparative ultrastructural analysis of the fungal endophytes in Cryptothallus mirabilis Malm. and other British thalloid hepatics. Journal of Bryology 13: 227-233..[Web of Science]
Puhlman J. Bucheli E. Swain M. J. Dunning N. Albersheim P. Darvill A. G. Hahn M. G.. 1994. Generation of monoclonal antibodies against plant cell wall polysaccharides. I. Characterization of a monoclonal antibody to a terminal -(12)-linked fucosyl-containing epitope. Plant Physiology 104: 699-710..[Abstract]
Qiu Y. L. Libo L. Wang B. Chen Z. Knoop V. Groth-Malonek M. Dombrowska O. Lee J. Kent L. Rest J. Estabrook G. F. Hendry T. A. Taylor D. W. Testa C. M. Ambros M. Crandall-Stotler B. Duff R. J. Stech M. Frey W. Quandt D. Davis C. C.. 2006. The deepest divergences in land plants inferred from phylogenomic evidence. Proceedings of the National Academy of Sciences, USA 103: 15511-15516..
Read D. J. Duckett J. G. Francis R. Ligrone R. Russell A.. 2000. Symbiotic fungal associations in "lower" land plants. Philosophical Transactions of the Royal Society London, B, Biological Sciences 355: 815-831..[CrossRef]
Renzaglia K. S. Schuette S. Duff R. J. Ligrone R. Shaw A. J. Mishler B. D. Duckett J. G.. 2007. Bryophyte phylogeny: advancing the molecular and morphological frontiers. Bryologist 110: 179-213..[CrossRef]
Rodriguez-Gálvez E. Mendgen K.. 1995. Cell wall synthesis in cotton roots after infection with Fusarium oxysporum. Differential deposition of callose, arabinogalactans, xyloglucans, and pectic components into walls, wall appositions, cell plates and plasmodesmata. Planta 197: 535-543..[Web of Science][Medline]
Russell J. Bulman S.. 2005. The liverwort Marchantia foliacea forms a specialized symbiosis with arbuscular mycorrhizal fungi in the genus Glomus. New Phytologist 165: 567-579..[CrossRef][Web of Science][Medline]
Saitou N. Nei M.. 1987. The neighbour-joining method: a new method for reconstructing phylogenetic trees. Molecular Biology and Evolution 4: 406-425..[Abstract]
Schmid E. Oberwinkler F.. 1993. Mycorrhiza-like interaction between the achlorophyllous gametophyte of Lycopodium clavatum L. and its fungal endophyte studies by light and electron microscopy. New Phytologist 124: 69-81..[CrossRef][Web of Science]
Schmid E. Oberwinkler F.. 1994. Light and electron microscopy of the host–fungus interaction in the achlorophyllous gametophyte of Botrychium lunaria. Canadian Journal of Botany 72: 182-188..
Schmid E. Oberwinkler F.. 1995. A light- and electron-microscopic study on a vesicular-arbuscular host–fungus interaction in gametophytes and young sporophytes of the Gleicheniaceae (Filicales). New Phytologist 129: 317-324..[CrossRef][Web of Science]
Schüßler A.. 2000. Glomus claroideum forms an arbuscular mycorrhiza-like symbiosis with the hornwort Anthoceros punctatus. Mycorrhiza 10: 15-21..[CrossRef][Web of Science]
Schüßler A. Mollenhauer D. Schnepf E. Kluge M.. 1994. Geosiphon pyriforme, an endosymbiotic association of fungus and cyanobacteria: the spore structure resembles that of arbuscular mycorrhizal (AM) fungi. Botanica Acta 107: 36-45..[Web of Science]
Schüßler A. Schwarzott D. Walker C.. 2001. A new fungal phylum, the Glomeromycota: phylogeny and evolution. Mycological Research 105: 1413-1421..[CrossRef][Web of Science]
Schwarzott D. Walker C. Schüßler A.. 2001. Glomus, the largest genus of the arbuscular mycorrhizal fungi (Glomales) is nonmonophyletic. Molecular Phylogenetics and Evolution 21: 190-197..[CrossRef][Web of Science][Medline]
Selosse M. A.. 2005. Are liverworts imitating mycorrhizas?. New Phytologist 165: 345-349..[CrossRef][Web of Science][Medline]
Selosse M. A. Le Tacon F.. 1998. The land flora: a phototrophfungus partnership?. Trends in Ecology and Evolution 13: 15-20..[CrossRef]
Simon L. Bousquet J. Lévesque R. C. Lalonde M.. 1993. Origin and diversification of endomycorrhizal fungi and coincidence with vascular land plants. Nature 363: 67-69..[CrossRef][Web of Science]
Smallwood M. Beven A. Donovan N. Neill S. J. Peart J. Roberts K. Knox J. P.. 1994. Localization of cell wall proteins in relation to the developmental anatomy of the carrot root apex. Plant Journal 5: 237-246..[CrossRef][Web of Science]
Smallwood M. Martin H. Knox J. P.. 1995. An epitope of rice threonine- and hydroxyproline-rich glycoprotein is common to cell wall and hydrophobic plasma membrane glycoproteins. Planta 196: 510-522..[Web of Science][Medline]
Smallwood M. Yates E. A. Willats W. G. T. Martin H. Knox J. P.. 1996. Immunochemical comparison of membrane-associated and secreted arabinogalactan-proteins in rice and carrot. Planta 198: 452-459..[CrossRef][Web of Science]
Smith F. A. Smith S. E.. 1997. Structural diversity in (vesicular)-arbuscular mycorrhizal symbioses. New Phytologist 137: 373-388..[CrossRef][Web of Science]
Smith S. E. Read D. J.. 1997. Mycorrhizal symbiosis. Academic Press, Cambridge, UK..
Steffan W. Kovàc P. Albersheim P. Darvill A. G. Hahn M. G.. 1995. Characterization of a monoclonal antibody that recognizes an arabinosylated (16)-ß-d-galactan epitope in plant complex carbohydrates. Carbohydrate Research 275: 295-307..[CrossRef][Web of Science][Medline]
Strullu D. G. Gourret J. P. Garrec J. P.. 1981. Microanalyse des granules vacuolaires des ectomycorhizes, endomycorhizes et endomycothalles. Physiologie Végétale 19: 367-378..[Web of Science]
Thompson J. D. Higgins D. G. Gibson T. J.. 1994. Clustal w: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position specific gap penalties and weight metrics choice. Nucleic Acids Research 22: 4673-4680..
Wang B. Qiu Y. L.. 2006. Phylogenetic distribution and evolution of mycorrhizas in land plants. Mycorrhiza 16: 299-363..[CrossRef][Web of Science][Medline]
Willats W. G. T. Limberg G. Buchholt H. C. van Alebeek G. Benen J. Christensen T. M. I. E. Visser J. Voragen A. Mikkelsen J. D. Knox J. P.. 2000. Analysis of pectic epitopes recognised by hybridoma and phage display monoclonal antibodies using defined oligosaccharides, polysaccharides, and enzymatic degradation. Carbohydrate Research 327: 309-320..[CrossRef][Web of Science][Medline]
Willats W. G. T. Marcus S. Knox J. P.. 1998. Generation of a monoclonal antibody specific to (15)--l-arabinan. Carbohydrate Research 308: 149-152..[CrossRef][Web of Science][Medline]
Willats W. G. T. Orfila C. Limberg G. Buchholt H. C. van Alebeck G. W. M. Voragen A. G. J. Marcus S. E. Christiensen T. M. I. E. Mikkelsen J. D. Murray B. S. Knox J. P.. 2001. Modulation of the degree and pattern of methyl-esterification of pectic homogalacturonan in plant cell walls. Journal of Biological Chemistry 276: 19404-19413..
CiteULike Complore Connotea Del.icio.us Digg Facebook Reddit Technorati Twitter What's this?
This article has been cited by other articles:
|
|
 |
|
  M. I. Bidartondo and J. G. Duckett Conservative ecological and evolutionary patterns in liverwort-fungal symbioses Proc R Soc B, February 7, 2010; 277(1680): 485 - 492. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. E. Graham, M. E. Cook, D. T. Hanson, K. B. Pigg, and J. M. Graham Structural, physiological, and stable carbon isotopic evidence that the enigmatic Paleozoic fossil Prototaxites formed from rolled liverwort mats Am. J. Botany, February 1, 2010; 97(2): 268 - 275. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Pressel, R. Ligrone, J. G. Duckett, and E. C. Davis A novel ascomycetous endophytic association in the rhizoids of the leafy liverwort family, Schistochilaceae (Jungermanniidae, Hepaticopsida) Am. J. Botany, May 1, 2008; 95(5): 531 - 541. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Z. Hock, P. Szovenyi, J. J. Schneller, Z. Toth, and E. Urmi Bryophyte diaspore bank: a genetic memory? Genetic structure and genetic diversity of surface populations and diaspore bank in the liverwort Mannia fragrans (Aytoniaceae) Am. J. Botany, May 1, 2008; 95(5): 542 - 548. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
