Michael Gillings and Andrew Holmes Department of Biological Sciences, Macquarie University, Sydney, NSW 2109, Australia
LONDON AND NEW YORK
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Plant Microbiology by Gillings, Michael.; Holmes, Andrew J. Publication: London, UK, New York, N.Y Taylor & Francis Routledge, 2004.
Chapter 1 The diversity, ecology and molecular detection of arbuscular mycorrhizal fungi
Rebecca Husband (1~18 页)
Plant Microbiology, Michael Gillings and Andrew Holmes
? 2004 Garland Science/BIOS Scientific Publishers, Abingdon. 1.1 Introduction
The vast majority of land plants rely on interactions with root symbionts to ensure adequate nutrient uptake. Of these root symbionts the most widespread and common are the arbuscular mycorrhizal (AM) fungi, which form associations with ca 60% of plant species (Smith and Read, 1997). The AM association is also arguably the most successful root symbiosis in evolutionary terms. Fossil evidence and molecular clock estimates indicate that the AM symbiosis originated at least 400 MYA (million years ago) and has not changed appreciably since (Simon et al., 1993a; Taylor et al., 1995). The association is almost universally distributed in early plant taxa with loss of the symbiosis only occurring more recently in ca 10% of plant families (Tester et al., 1987; Trappe, 1987). It is therefore hypothesised that the AM symbiosis was instrumental in ensuring the successful colonisation of land by plants (Simon et al., 1993a). The AM symbiosis is widely accepted to be mutualistic. The most obvious benefit to the fungus is a ready supply of carbon, whilst the plant gains access to nutrients (most notably phosphorus) that might not be available from root uptake alone (Smith and Read, 1997). Other benefits to the plant include improved water relations and protection against pathogens (Newsham et al., 1995). Plants associated with AM fungi often exhibit increased growth and survival (Smith and Read, 1997) although critically the level of benefit depends on a variety of factors including the particular host-fungal combination (Helgason et al., 2002; Streitwolf-Engel et al., 1997; van der Heijden et al., 1998a). The differential effects individual AM fungi have on plant performance are therefore proposed to have a major influence on ecosystem functioning, from affecting productivity (Klironomos et al.,
2000), to altering competitive interactions (Gange et al., 1993; Grime et al., 1987; Hartnett and Wilson, 1999; O’ Connor et al., 2002), and influencing the overall diversity of plant communities (van der Heijden et al., 1998b). Yet despite the important role AM fungi play in ecosystem functioning, little is known of the community structure and ecology of the fungi themselves. To date, fewer than 200 AM fungal species have been identified (Morton and Benny, 1990). This apparent low global diversity of AM fungi compared to their associated host plant communities has led to the widespread belief, only now being challenged, that AM fungi are a functionally homogeneous group (Smith and Read, 1997). Indeed, with few exceptions, the majority of ecological studies have ignored the functional diversity of individual AM fungi and grouped all the fungi into a single class. To be fair, this is because AM fungi are notoriously difficult to study. Firstly, AM fungi are obligate biotrophs and we have yet to find a way of culturing them independently of their plant hosts. Secondly, AM fungi exhibit a very low morphological diversity, making the reliable identification of different species difficult. Fungal structures formed internally within the host root possess few diagnostic characters; therefore the taxonomy of AM fungi is based on the subcellular structures of asexual spores. The spores of AM fungi are relatively large, easy to extract from the soil and have enough characteristics to enable identification to species level by experienced personnel (see the International Culture Collection of Arbuscular and Vesicular Mycorrhizal Fungi (INVAM) website http://invam.caf.wvu.edu for more information). Estimating the diversity of AM fungi in the field has therefore traditionally relied on detecting the spores present in the soil. Since the spores tend to be ephemeral a complementary approach involves growing ‘ trap’ plants in field soils in the greenhouse and analysing the spores produced. Unfortunately, these methods are problematic because fungal sporulation rates are influenced both by the environment and the host plant species (Bever et al., 1996; Eom et al., 2000; Morton et al., 1995), therefore spore counts are not a direct measure of diversity. Furthermore, the population of spores in the soil may bear little relation to the AM fungal populations colonising roots (Clapp et al., 1995). One of the most important methodological advances in the study of AM communities has been the application of the polymerase chain reaction (PCR) to directly identify the AM fungi in planta. Recovering sequence information from the field gives us direct access, without relying on culturing, to the AM fungi present in roots, the ecologically significant niche. Sequence information also provides us with the means to consistently distinguish between morphologically similar taxa. Thus molecular tools now present us with new opportunities for understanding the role of AM fungi in the environment. In this chapter some of the methods
available to study AM fungal diversity in the field are outlined and the key findings from such studies are reviewed.
1.1.1 The taxonomy of AM fungi
Based on morphological characters, fewer than 200 AM fungal species have been described (Morton and Benny, 1990). They can be divided into five families, the Acaulosporaceae (Acaulospora, Entrophospora), Gigasporaceae (Gigaspora, Scutellospora) and Glomaceae (Glomus) (Morton and Benny, 1990) plus the newly described lineages (Archaeosporaceae) and Paraglomaceae (Paraglomus) (Morton and Redecker, 2001). Traditionally these families have been placed in the order Glomales, phylum Zygomycota, but in recent years both morphological and molecular evidence have indicated that the phylum Zygomycota as currently defined cannot be sustained. Firstly, many of the organisms assigned to it, including AM fungi, are not known to have a sexual stage, i.e. they do not form zygosporangia (Benny, 1995). Secondly, phylogenetic analyses based on sequence data have demonstrated that the lineages ascribed to the Zygomycota do not share a common ancestor, i.e. the phylum Zygomycota is polyphyletic (O’ Donnell et al., 2001; Tehler et al., 2000). Consequently, based on a small subunit (SSU) ribosomal gene (rDNA) phylogeny, Schü ler et al. (2001b) have proposed a new classification for the AM fungi, ? removing them from the Zygomycota and placing them in a new phylum the Glomeromycota. The analyses of Schü ler et al. (2001b) indicate that the AM fungi can be separated into a monophyletic clade that is not closely ? related to any of the Zygomycota lineages, but is instead probably diverged from the same common ancestor as the Ascomycota and Basidiomycota. At the lower taxonomic level, the SSU rDNA phylogeny indicates a large genetic diversity within the genus Glomus that is not reflected by any of the morphological characters available (Schwarzott et al., 2001). Sequences for the AM fungi within this genus can have genetic distances as large as that between the families Acaulosporaceae and Gigasporaceae and as a result the classification of Schü ler et al. (2001b) also proposes a ? new family and order ranking (see Figure 1.1). The proposed order Diversisporales contains the two traditional family groupings of the Acaulosporaceae and Gigasporaceae, plus a new family Diversisporaceae fam. ined. consisting of some of the AM fungi originally classified within the genus Glomus. The remaining ‘ classical’ Glomus spp. have been placed in the order Glomerales, which clearly separates into two distinct family-ranked clades, Glomus Group A and B. Two further orders are proposed, the Paraglomerales (single family Paraglomeraceae) and the Archaeosporales (two families, Archaeosporaceae and Geosiphonaceae). As currently defined, the family Archaeosporaceae is paraphyletic. The type species for the Geosiphonaceae is a
non-mycorrhizal fungus, Geosiphon pyriforme, which forms an association with cyanobacteria. Previously it had been proposed that Geosiphon represented the ancestral precursor to AM fungi (Gehrig et al., 1996), but the recent rDNA data reveal it is in fact closely related to the Archaeosporaceae and consequently the phylum Glomeromycota, as defined, includes both mycorrhizal and non-mycorrhizal fungi. The classification of Schü ler et al. (2001b) will necessarily evolve as more information becomes available. ? The lack of convincing morphological characters for many of the groupings and the presence of multiple sequence variants within single spores of AM fungi (discussed in Section 1.2), means that more detailed work is needed before the taxonomy and systematics of AM fungi are truly understood. Nonetheless the classification of Schü ler ? et al. (2001b) represents the basis for a new taxonomy for AM fungi, finally acknowledging what has long been obvious, namely that AM fungi are not typical Zygomycetes
Figure 1.1. Phylogeny of the Glomeromycota (neighbour-joining analysis of SSU rDNA sequences) indicating the families and orders proposed by Schü ler et al., (2001b). Bootstrap values >70% (1000) ? replicates are shown. Asterisks (*) identify the AM fungal taxa previously classified within the genus Glomus. A putative choanozoan, C. limacisporum L42528, (Cavalier-Smith and Allsopp, 1996) is used as an outgroup. Accession numbers are as follows: Ar. trappei Y17634, Ge. pyriforme AJ276074, Ar. leptoticha AJ301861, P. occultum AJ276082, P. brasilianum AJ301862, G. υ iscosum Y17652, G. etunicatum Y17639, G. luteum AJ276089, G. lamellosum AJ276087, G. mosseae AJ418853, G. sinuosum AJ133706, G. intraradicesAJ301859 G. clarum AJ276084, E. colambiana Z14006, A. spinosa Z14004, A. scrobiculata AJ306442, A. longula AJ306439, G. etunicatum AJ301860, G. versiforme X86687, G. spurcum Y17650, S. dipapillosa Z14013, S. cerradensis AB041344, S. callospora AJ306443, S. projecturata AJ242729, Gi. albida Z14009, Gi. margarita X58726, Gi. Gigantea Z14010.
1.2 Molecular techniques used to study AM fungi in the field
Molecular techniques have the potential to revolutionise the study of AM fungi in the environment. Previously we have only had access to those AM fungi amenable to trap culture or actively sporulating in the field. Now a variety of nucleic acid-based strategies have been developed that enable the AM fungi to be characterised independently of spore formation. The majority of sequence information for AM fungi is derived from the ribosomal RNA genes (rDNA). In most organisms the ribosomal RNA genes are present in multiple copies arranged in tandem arrays. Each repeat unit consists of genes encoding a small (SSU or 18S) and a large (LSU or 28S) subunit, separated by an internal transcribed spacer (ITS), which includes the 5.8S rRNA gene (Figure 1.2). The SSU and 5.8S genes evolve relatively slowly and are useful for studies of distantly related organisms. The LSU and ITS regions evolve more quickly and are useful for fine-scale differentiation between species. In most organisms, the process of concerted evolution ensures that within an individual the multiple copies of rDNA are identical, but early studies looking at the genetic diversity of AM fungi revealed an unexpectedly high degree of ITS sequence variation within single spores (Lloyd-MacGilp et al., 1996; Sanders et al., 1995). Sequence divergence within single spores has since been detected in the ITS of many different species (Antoniolli et al., 2000; Jansa et al., 2002; Lanfranco et al., 1999; Pringle et al., 2000). The diversity is not restricted to the ITS, but has also been detected in the SSU (Clapp et al., 1999) and extensive intrasporal diversity appears to exist in the LSU (Clapp et al., 2001; Rodriguez et al., 2001). Most recently, intrasporal variation has been detected in a gene encoding a binding protein (BiP) (Kuhn et al., 2001). Currently the full implications of the intrasporal variation present within the Glomeromycota are unclear. One obvious problem is that there is no straight-forward correlation between sequence identity and species identity, and as a consequence there is no phylogenetic species concept for AM fungi. Furthermore there is no straightforward correlation between the number of sequence variants detected within a root and the number of separate infection events. Thus, ecological studies that use molecular markers to study AM fungal diversity are limited as to the conclusions they can make. Not only is it impossible to ascribe a species name to a sequence type, but it is also impossible to determine how many different AM fungi are colonising each root. As a consequence the majority of studies reviewed in this chapter have simply placed the AM fungi into groups based Currently the full implications of the intrasporal variation present within the Glomeromycota are unclear. One obvious problem is that there is no straight-forward correlation between sequence identity and species identity, and as a consequence there is no phylogenetic species concept for AM fungi. Furthermore there is no straightforward
correlation between the number of sequence variants detected within a root and the number of separate infection events. Thus, ecological studies that use molecular markers to study AM fungal diversity are limited as to the conclusions they can make. Not only is it impossible to ascribe a species name to a sequence type, but it is also impossible to determine how many different AM fungi are colonising each root. As a consequence the majority of studies reviewed in this chapter have simply placed the AM fungi into groups based
Figure 1.2. A repeat unit of the fungal ribosomal RNA genes, showing the position of primers commonly used for the study of AM fungi.
1.2.1 Community detection
One of the most common goals in arbuscular mycorrhizal research is to determine the role AM fungi play in ecosystem functioning. Previously any such research has been severely limited by the lack of basic information such as the identities and distributions of the fungi. Whilst molecular techniques can help us gain such information, the methods themselves are limited by the available molecular markers. As a rule, in the wider field of molecular ecology, a single set of ribosomal primers is quickly established that is sufficient for all preliminary investigations of the organism(s) in question (see Avise, 1994; Carvalho, 1998). Not so the arbuscular mycorrhizas; as yet there is no single method that can reliably measure in planta diversity. Separate from the issue of intraspecies diversity, we have yet to develop markers that can differentiate all AM fungal sequences from non-AM fungal and plant sequences. With the discovery of the highly divergent families of the Archaeosporaceae and Paraglomeraceae the sequence divergence within the Glomeromycota is considerable, so it may never be possible to rely on a single molecular marker. Currently, a compromise must be made between the level of genetic resolution and the number of lineages detected. The first paper to apply molecular techniques to AM research appeared 10 years ago (Simon et al., 1992). The authors used general eukaryotic primers to amplify SSU sequences from spores and, based on this information, designed the primer VANS1 which they hoped to be a general AM fungal primer. They subsequently designed family-specific primers (VAGIGA, VAGLO, VAACAU) which, when teamed with VANS1, enabled the direct
amplification of AM fungi from within plant roots (Simon et al., 1993b). Although these primers appeared to work well on roots from microcosm studies, in the field they were more problematic (Clapp et al., 1995). It was later revealed that the VANS1 site is not well conserved throughout the Glomeromycota (Clapp et al., 1999; Schü ler ? et al., 2001a). Helgason et al. (1998) also targeted the SSU when they designed the AM1 primer to exclude plant sequences and preferentially amplify AM fungal sequences. Coupled with a general eukaryotic primer, it has been successfully utilised in the field to determine the diversity of AM fungi from many different habitats (Daniell et al., 2001; Helgason et al., 1998, 1999, 2002; Husband et al., 2002a, b; Kowalchuk et al., 2002; Vandenkoornhuyse et al., 2002). However, new sequence data have revealed that the AM1 primer is not well conserved in certain divergent lineages, the Archaeosporaceae and the Paraglomeraceae (Morton and Redecker, 2001). The AM1 primer also contains two mis-matches for sequences belonging to the Glomus group B clade defined by Schü ler ? et al. (2001b). In addition, in some habitat types, relatively high proportions (up to 30%) of non-AM fungi, mainly pyrenomycetes, are co-amplified (Daniell et al., 2001). Even so, at this present time AM1 remains the most broadly applicable single primer suitable for field studies, reliably detecting the three traditional families, and having numerous studies that provide useful comparisons. In contrast, Kj? 1ler and Rosendahl, (2001) have designed LSU primers specific to a subgroup in the Glomeraceae. The primers LSURK4f and LSURK7r are used in a nested PCR following amplification with general eukaryotic primers (LSU0061 and LSU0599; van Tuinen et al., 1998) and are designed to amplify a lineage within the Glomus group A clade, including G.mosseae, G.caledonium and G.geosporum. The authors took this approach because preliminary characterisation of the spore populations in their field site determined that the AM fungal community was dominated by many very closely related Glomus species that would be very difficult to distinguish using the SSU gene. Therefore they utilised the higher diversity of the LSU to separate the different species, enabling community comparisons to be made within this subgroup. A series of group-specific primers targeting the major lineages within the Glomeromycota has also been designed by Redecker (2000). These primers amplify parts of the SSU, the ITS and the 5.8S gene, although they have yet to be used in the field. In the field, most plant roots are colonised by more than one AM type; therefore, unless single-taxon primers are used, a way must be found to separate the different types. The majority of the studies using the AM1 primer have used an approach based on cloning, PCR, and restriction fragment length polymorphism (RFLP) to divide the AM fungi
into classes. Examples of each class can then be sequenced to give an insight into their identity. If it is assumed each fungal type is amplified and cloned proportionally, then the numbers of each class can be perceived as an approximate estimate of their proportion in the root. Although yielding much valuable information, the cloning step is both expensive and labour-intensive. Recently, Kowalchuk et al. (2002) adopted a different technique, denaturing gradient gel electrophoresis (DGGE), Muyzer et al., 1993), to characterise the AM communities. Separation of the different AM types depends on the melting behaviour of the DNA sequence and, in theory, DGGE is sensitive enough to detect differences of a single base pair. A slightly different gel-based technique, single-strand conformation polymorphism (SSCP, Orita et al., 1989) was used by Simon et al. (1993b) and Kj? 1ler and Rosendahl (2001). The PCR product is denatured immediately before loading on a non-denaturing gel, and separation is achieved through migrational differences between the sequences as they adopt different conformations within the gel. An alternative approach, terminal restriction fragment lengthpolymorphism (T-RFLP; Liu et al., 1997) was used by Vandenkoornhuyse et al. (personal communication). This method uses a PCR in which the primers are fluorescently labelled. After amplification, the PCR product is digested with one or more enzymes generating terminal-labelled fragments that are characteristic in size. In theory, with the appropriate combination of genetic marker and restriction enzyme, terminal fragments can be generated that are diagnostic of individual species.
1.2.2 Specific detection
Frequently it is desirable to focus on the ecology of specific fungal isolates. The method devised by van Tuinen et al. (1998) provides a good example of how species-specific primers can track different AM fungal strains in mixed inoculum experiments. The authors designed primers specific for the LSU of each inoculant species, using them in a second round of amplification to gain information on the competitive interactions of the various isolates. This approach has successfully been used and expanded in microcosm experiments testing the effect of sewage sludge treatments on AM fungi (Jacquot et al., 2000; Jacquot-Plumey et al., 2001) and directly in the field studying the effect of heavy-metal polluted soils (Turnau et al., 2001). The study by Turnau et al. (2001) further serves to illustrate the discrepancy between spore and root populations of AM fungi. Even though the authors characterised the spore population at their field site and designed primers for all the species isolated, many plant roots did not yield amplified sequences despite being clearly colonised. The main utility of molecular markers able to differentiate between closely related strains is in the field of molecular taxonomy. The ITS region has been used extensively for such studies and the universal primers ITS1
and ITS4, designed by White et al. (1990), have proved especially useful. However, due to the sequence variation present within single spores of AM fungi, the ITS region cannot be used as a taxonomic tool as it has been in other organisms. Nonetheless, the ITS1 and ITS4 primers have been used extensively in AM research to study the nature of this intrasporal sequence variation itself (Antoniolli et al., 2000; Jansa et al., 2002; Lanfranco et al., 1999; Lloyd-MacGilp et al., 1996; Pringle et al., 2000; Sanders et al., 1995).
1.3 The molecular diversity of AM fungi colonising roots in the field
At present we do not know what level of genetic diversity is meaningful in an ecological context, therefore it is not possible to fully interpret the results of ecological studies that use molecular markers. Despite this limitation, molecular techniques have so far provided much valuable information on the diversity of AM fungi across a variety of habitats. The first study (Clapp et al., 1995) that used molecular techniques to analyse the diversity of AM fungi colonising roots in the field used the family-specific primers designed by Simon et al. (1992, 1993b). The authors compared molecular data for the presence or absence of each of the families in roots, with counts of spores isolated from the surrounding soil. The morphological and molecular data were largely in agreement for Acaulospora and Scutellospora types, but there was a large discrepancy for Glomus types, whereby Glomus spores were rarely found in the soil yet Glomus types were frequent colonisers of the roots (Clapp et al., 1995). Thus this study showed conclusively what had long been suspected; spore populations do not accurately reflect the AM fungi colonising roots. To date the most extensive molecular investigations have all used the AM1 primer, making it possible to draw comparisons between the AM communities in a seminatural woodland (Helgason et al., 1999, 2002), arable sites (Daniell et al., 2001), a seminatural grassland (Vandenkoornhuyse et al., 2002), coastal sand dunes (Kowalchuk et al., 2002) and a tropical forest (Husband et al., 2002a, b). A summary of the levels of AM fungal diversity detected within these habitats is given in Table 1.1. Although different degrees of sampling intensity make it difficult to make direct comparisons, collectively these studies appear to reveal an approximate correlation between above- and below-ground diversity. In itself this approximate correlation has an important implication. Van der Heijden et al. (1998b) found that plant diversity increased with increasing AM fungal diversity in their experimental microcosm system. They suggested that the increase in plant diversity resulted from the growth of different plants being stimulated by different fungal species and consequently that AM fungal identity and diversity were potential determinants of plant community structure. Although no causal relationship can be drawn
from the molecular field data, the demonstration that in the field there is a link between plant and AM fungal diversity is consistent with the hypothesis that AM fungi are potential determinants of ecosystem diversity. These molecular data also reveal large differences in the AM community composition between the different habitat types. The arable sites, seminatural grassland and tropical forest are all heavily dominated by Glomus types, both in terms of the number of types and their abundance. In contrast, the seminatural woodland AM community is more evenly distributed between Acaulospora and Glomus types, though in terms of abundance, one of the woodland hosts Hyacinthoides non-scripta (bluebell), is heavily dominated by a Scutellospora type early in the growing season (Helgason et al., 1999). No Acaulospora types were detected in the AM community colonising Ammophila arenaria in coastal sand dunes; instead the community contained equal numbers of Glomus and Scutellospora types (Kowalchuk et al., 2002). Where the experimental design allows it, extensive spatial and temporal heterogeneity is revealed within each habitat. Kowalchuk et al. (2002) were able to detect clear differences between the AM communities colomsing Ammophila in vital and degenerating stands. Not only were the degenerating stands depauperate relative to the vital stands, but the relative signal intensities of the samples from the degenerating stands tended to be substantially reduced. The AM community colonising bluebells in the seminatural woodland shows a clear seasonal succession, initially being dominated by a Scutellospora type which later in the season gives way to Glomus types if the dominant canopy is Acer pseudoplatanus, or Acaulospora types if the dominant canopy is Quercus petraea (Helgason et al., 1999). These trends match well with the data from morphological analyses of the fungi colonising bluebell roots (Merryweather and Fitter, 1998a, b). Similarly, Husband et al. (2002a, b) detected a replacement over time in the presence and abundance of AM fungi colonising cohorts of seedlings in a tropical forest. The grassland mycorrhizal community was also shown to change at each sampling period, and the authors suggested that a shift in field management from grazing to mowing and the subsequent decrease in organicmatter might be responsible (Vandenkoornhuyse et al., 2002).
Table 1.1. A summary of the levels of AM fungal diversity detected across
Sand dunea Arableb Woodlandc Grasslandd Tropical foreste
No. of host No. of species roots
1 4 6 2 2 / 79 71 49 48
No. of No. of No. of No. of Total no. clones Acualo-spora sp. Glomus sp. Giga-spora sp. of types
n.a. 303 257 2001 1383 0 1 5 2 1 3 6 6 15 21 3 1 1 1 1 6 8 13* 18 23
Data from aKowlachuk et al., (2002); bDaniell et al., (2001); cHelgason et al., (1999, 2002); Vandenkoornhuyse et al., (2002); and eHusband et al., (2002a). *An Archaeospora type was also detected.
At different spatial scales, the woodland, grassland and tropical forest studies, all detected non-random associations between the plant community and the AM fungal community. The woodland AM community was influenced by the dominant canopy type (Helgason et al., 1999); whereas the grassland AM community (at a single site) was shown to be significantly different between the two host species, Agrostis capillaris and Trifolium repens (Vandenkoornhuyse et al., 2002). The tropical AM community was also signiflcantly different between the host species, but the environment was found to have a greater influence, such that differences between host species were site-specific (Husband et al., 2002a). Again, although no causal relationship can be made, these non-random patterns of association have an important implication. In recent years, various microcosm studies have shown that the AM fungal community can affect plant diversity or υ υ ice ersa (Bever, 2002; Bever, et al., 1996; Burrows and Pfleger, 2002; Eom et al., 2000; Helgason et al., 2002; Sanders and Fitter, 1992; van der Heijden et al., 1998 a, b). If such processes occur in the environment, some degree of host preference in natural populations is highly likely. Indeed, by selecting morphologically distinct fungal species, McGonigle and Fitter (1990) were the first to demonstrate that non-random associations between different hosts and AM fungi exist in the field. These molecular data support the findings of McGonigle and Fitter (1990) and suggest that in the environment AM fungi may commonly exhibit a host preference. The study by Helgason et al. (2002) is especially relevant because not only did they demonstrate that root colonisation, symbiont compatibility and plant performance varied with each fungus-plant combination in the greenhouse, but they were able to link the functioning of the mycorrhizae with the patterns of association between plants and fungi found in the fleld. For example, the authors suggested that one of the fungal types, Glomus sp.
UY1225, appears to be a ‘ typical’AM fungus. In the field it shows a relatively broad host range and in the laboratory study Glomus sp. UY1225 provided some benefit to most of the plant species without greatly benefiting any of them. The only plant species it did not colonise extensively in pots was Acer, the only species in the field survey from which it was absent. In fact, the only fungus to colonise and benefit Acer in the laboratory study was Glomus hoi UY110. In the field the AM type Glo9, which is very closely related to, but distinct from, G. hoi, is found almost exclusively in Acer roots. The authors suggest that the sequence variation detected within G. hoi might represent a fraction of the variation within a single species that would in fact include the field-derived Glo9 sequences. Unfortunately, the low clone numbers generated from the field study make it impossible to test this idea. Even so, the authors argue that given the large impact of G. hoi on Acer growth, it would be very unexpected to find two functionally unrelated taxa restricted to the roots of Acer in the field. If, in the future, it is demonstrated that G. hoi and Glo9 are one and the same, the study by Helgason et al. (2002) will have provided the first ever evidence of functional selectivity within the arbuscular mycorrhizal symbiosis. The AM types recognised by these molecular techniques cannot be equated directly with the formal species that are identified on the basis of spore morphology. Even so, many of the sequence types have been recovered repeatedly in different studies, and there are examples of identical sequences being isolated from different habitats (see Figure 1.3 and * in Figure 1.4). It would appear that many of these types represent entities as widespread and stable as those defined by morphology. Furthermore, the SSU rDNA region amplified by the NS31/AM1 primers appears to provide a level of discrimination at approximately the species level (Figure 1.3). For example, the G. mosseae clade contains numerous sequences from different spores and cultures revealing a level of intraspecies variation comparable to the variation in the field-derived sequence type Glolb. Similarly, the clade containing the sequence type Glo3 contains numerous sequences that have been isolated repeatedly both from different hosts and time points within a habitat, and from different habitats. A sequence from the Glomus sp. isolate UY1225 trapped from the woodland soil (Helgason et al., 2002) matches many of the field-derived Glo3sequences. Overall this clade reveals a similar level of variation as the G. mosseae/Glo1b clade. In contrast the sequence type Glo8 contains three distinct groups, one of which includes sequences from G. fasiculatum and G. vesiculiferum, another containing a G. fasiculatum sequence and the third containing sequences from G. intraradicies. There are too few culture-derived sequences to make many comparisons of this nature, and it must be acknowledged that if more sequences per isolate per spore were characterised, the level of intraspecies variation within the SSU could turn out to be much greater than presently recognised. However, the study by Kowalchuk et al. (2002) used DGGE to
characterise various AM fungal isolates and detected no intraspecies variation, with the exception of G. clarum that consistently yielded two bands. Critically, they analysed both single-spore and multispore extracts thus maximising the probability of detecting intraspecies variation if it existed. In theory DGGE is sensitive enough to detect differences of a single basepair, but Kowalchuk et al. (2002) were not able to distinguish between two closely related species Gi. margarita and Gi. albida, the sequences for which differ by approximately five basepairs. Therefore, based on the data currently available, it would seem that small levels of intraspecies variation are present in this region of the SSU, but the variation is not so great as to be prohibitive to community studies. The phylogenetic tree shown in Figure 1.4 contains a single example of the different Glomus group A sequences isolated from the various habitats. As can be seen, with the exception of the Glo types already discussed, very few of the field-derived sequences group with sequences from AM fungi in culture. This phenomenon is not restricted to studies using the AM1 primer. Kj? 1ler and Rosendahl (2001) deliberately designed their LSU primers to focus on a single lineage that includes G. mosseae, G. claroideum and G. geosporum, because these were the species that had been isolated from their field site. Yet despite limiting their study to these groups, they too recovered very few field sequences that matched known isolates. The simplest explanation for these observations is that the number of AM fungi in culture represent but a fraction of the true AM fungal diversity. Recently both Bever et al. (2001) and Helgason et al. (2002) have put forward arguments to this effect. Helgason et al. (2002) also challenge the traditional assumption that AM fungi are not host-specific. They argue such an assumption is based on the fact that: (i) fewer than 200 species have been described; and (ii) the AM fungi in culture tend to have a broad host range. However, they suggest there could be large numbers of as yet undescribed AM fungi that we have not managed to culture precisely because they are more host-selective. The growing number of non-random associations detected between different AM fungi and hosts in the fleld, plus the minimal overlap between sequences derived from the field and from cultured isolates, would seem to support their claims.
Figure 1.3. Neighbour-joining phylogenetic tree of the Glolb, Glo3 and Glo8 field-derived sequences recovered from W seminatural woodland (Helagson et al., 1999, 2002); A, arable sites (Daniell et al., 2001); G, seminatural grassland (Vandenkoornhyuse et al., 2002) and T, tropical forest (Husband et al., 2002a, b). Bootstrap values >70% are shown (1000 replicates). Multiple identical sequences of the number indicated have been recovered.
Figure 1.4. Neigbour-joining phylogenetic tree showing examples of the different Glomus-group A sequences isolated from W seminatural woodland (Helagson et al., 1999, 2002); A, arable sites (Daniell et al., 2001); G, seminatural grassland (Vandenkoornhyuse et al., 2002) and T, tropical forest (Husband et al., 2002a, b). Bootstrap values >70% are shown (1000 replicates). Asterisks (*) identify identical sequences recovered from different habitats. Brackets indicate identical sequence types recovered from different habitats.
The application of molecular techniques to the ecological study of AM fungi has led to a number of valuable insights. It has repeatedly been demonstrated that the AM community composition within roots is diverse, changes radically between different habitats, and within habitats between different time points and plant species. This variation itself is proof that the AM fungi in the field are not ecologically equivalent. However, the ecological role of AM fungi will never be fully appreciated until we understand the relationship between morphological, functional and molecular diversity. The challenge for the future is to resolve the genetic structuring of AM fungi so that we may address ecological questions in a determined manner and ultimately establish the link between above- and below-ground ecosystem diversity.
Antoniolli, Z.I., Schachtman, D.P., Ophel-Keller, K., Smith, S.E. (2000) Variation in rDNA ITS sequences in Glomus mosseae and Gigaspora margarita spores from a permanent pasture. Mycol. Res. 104:708– 715. Avise, J.C. (1994) Molecular Markers, Natural History and Evolution. London: Chapman and Hall. Benny, G.L. (1995) Classical morphology in Zygomycete taxonomy. Can. J. Bot.-Reυ Can. Botan. 73: . S725– S730. Bever, J.D. (2002) Host-specificity of AM fungal population growth rates can generate feedback on plant growth.Plant Soil 244:281– 290. Bever, J.D., Morton, J.B., Antonovics, J., Schultz, P.A. (1996) Host-dependent sporulation and species diversity of arbuscular mycorrhizal fungi in a mown grassland. J. Ecol. 84:71– 82. Bever, J.D., Schultz, P.A., Pringle, A., Morton, J.B. (2001) Arbuscular mycorrhizal fungi: More diverse than meets the eye, and the ecological tale of why. Bioscience 51:923– 931. Burrows, R.L., Pfleger, F.L. (2002) Arbuscular mycorrhizal fungi respond to increasing plant diversity. Can. J. Bot.-Reυ Can. Botan. 80:120– . 130. Carvalho, G.R. (1998) Advances in Molecular Ecology. IOS Press, Oxford. Cavalier-Smith, T., Allsopp, M. (1996) Corallochytrium, an enigmatic non-flagellate protozoan related to choanoflagellates. Euro. J. Protistol. 32:306– 310. Clapp, J.P., Young, J.P.W., Merryweather, J.W., Fitter, A.H. (1995) Diversity of fungal symbionts in arbuscular mycorrhizas from a natural community. New Phytol. 130:259– 265. Clapp, J.P., Fitter, A.H., Young, J.P.W. (1999) Ribosomal small subunit sequence variation within spores of an arbuscular mycorrhizal fungus, Scutellospora sp. Mol. Ecol. 8:915– 921. Clapp, J.P., Rodriguez, A., Dodd, J.C. (2001) Inter- and intra-isolate rRNA large subunit variation in Glomus coronatum spores. New Phytol. 149:539– 554. Clapp, J.P., Helgason, T., Daniell, T.J., Young, J.P.W. (2002) Genetic studies of the structure and diversity of arbuscular mycorrhizal communities. In: van der Heijden, M.G.A, Sanders, I.R. (eds) Mycorrhizal Ecology, pp. 201– 224. Heidelberg: Springer.
Daniell, T.J., Husband, R., Fitter, A.H., Young, J.P.W. (2001) Molecular diversity of arbuscular mycorrhizal fungi colonising arable crops. FEMS Microbiol. Ecol. 36:203– 209. Eom, A.H., Hartnett, D.C., Wilson, G.W.T. (2000) Host plant species effects on arbuscular mycorrhizal fungal communities in tallgrass prairie. Oecologia 122:435– 444. Gange, A.C., Brown, V.K., Sinclair, G.S. (1993) Vesicular-arbuscular mycorrhiza fungi— a determinant of plant community structure in early succession. Function. Ecol. 7:616– 622. Gehrig, H., Schü ler, A., Kluge, M. (1996) Geosiphon pyriforme, a fungus forming endocytobiosis with ? Nostoc (Cyanobacteria), is an ancestral member of the Glomales: Evidence by SSU rRNA analysis. J. Mol. Eυ 43:71– ol. 81. Grime, J.P., Mackey, J.M.L., Hillier, S.H., Read, D.J. (1987) Floristic diversity in a model system using experimental microcosms. Nature 328:420– 422. Hartnett, D.C., Wilson, G.W.T. (1999) Mycorrhizae influence plant community structure and diversity in tallgrass prairie. Ecology 80:1187– 1195. Helgason, T., Daniell, T.J., Husband, R., Fitter, A.H., Young, J.P.W. (1998) Ploughing up the wood-wide web? Nature 394:431. Helgason, T., Fitter, A.H., Young, J.P.W. (1999) Molecular diversity of arbuscular mycorrhizal fungi colonising Hyacinthoides non-scripta (bluebell) in a seminatural woodland. Mol. Ecol. 8:659– 666. Helgason, T., Merryweather, J.W., Denison, J., Wilson, P., Young, J.P.W., Fitter, A.H. (2002) Selectivity and functional diversity in arbuscular mycorrhizas of co-occurring fungi and plants from a temperate deciduous woodland. J. Ecol. 90:371– 384. Husband, R., Herre, E.A., Turner, S.L., Gallery, R., Young, J.P.W. (2002a) Molecular diversity of arbuscular mycorrhizal fungi and patterns of host association over time and space in a tropical forest.Mol. Ecol. 11:2669– 2678. Husband, R., Herre, E.A., Young, J.P.W. (2002b) Temporal variation in the arbuscular mycorrhizal communities colonising seedlings in a tropical forest. Fems Microbiol. Ecol. 42:131– 136. Jacquot, E., van Tuinen, D., Gianinazzi, S., Gianinazzi-Pearson, V. (2000) Monitoring species of arbuscular mycorrhizal fungi in planta and in soil by nested PCR: application to the study of the impact of sewage sludge. Plant Soil 226:179– 188. Jacquot-Plumey, E., van Tuinen, D., Chatagnier, O., Gianinazzi, S., Gianinazzi-Pearson, V. (2001) 25S rDNA-based molecular monitoring of glomalean fungi in sewage sludge-treated field plots. Enυ iron. Microbiol. 3:525– 531. Jansa, J., Mozafar, A., Banke, S., McDonald, B.A., Frossard, E. (2002) Intra- and intersporal diversity of ITS rDNA sequences in Glomus intraradices assessed by cloning and sequencing, and by SSCP analysis. Mycol. Res. 106:670– 681. Kj? ller, R., Rosendahl, S. (2001) Molecular diversity of glomalean (arbuscular mycorrhizal) fungi determined as distinct Glomus specific DNA sequences from roots of field grown peas. Mycol. Res . 105:1027– 1032. Klironomos, J.N., McCune, J., Hart, M., Neville, J. (2000) The influence of arbuscular mycorrhizae on the relationship between plant diversity and productivity. Ecol. Lett. 3:137– 141. Kowalchuk, G.A., De Souza, F.A., Van Veen, J.A. (2002) Community analysis of arbuscular mycorrhizal fungi associated with Ammophila arenaria in Dutch coastal sand dunes. Mol. Ecol. 11:571– 581. Kuhn, G., Hijri, M., Sanders, I.R. (2001) Evidence for the evolution of multiple genomes in arbuscular mycorrhizal fungi. Nature 414:745– 748.
Lanfranco, L., Delpero, M., Bonfante, P. (1999) Intrasporal variability of ribosomal sequences in the endomycorrhizal fungus Gigaspora margarita. Mol. Ecol. 8:37– 45. Liu, W.T., Marsh, T.L., Cheng, H., Forney, L.J. (1997) Characterization of microbial diversity by determining terminal restriction fragment length polymorphisms of genes encoding 16S rRNA. Appl. Enυ iron. Microbiol. 63:4516– 4522. Lloyd-MacGilp, S.A., Chambers, S.M., Dodd, J.C., Fitter, A.H., Walker, C., Young, J.P.W. (1996) Diversity of the ribosomal internal transcribed spacers within and among isolates of Glomus mosseae and related mycorrhizal fungi. New Phytol. 133:103– 111. McGonigle, T.P., Fitter, A.H. (1990) Ecological specificity of vesicular arbuscular mycorrhizal associations. Mycol. Res. 94:120– 122. Merryweather, J., Fitter, A. (1998a) The arbuscular mycorrhizal fungi of Hyacinthoides non-scripta— I. Diversity of fungal taxa. New Phytol. 138:117– 129. Merryweather, J., Fitter, A. (1998b) The arbuscular mycorrhizal fungi of Hyacinthoides non-scripta— II. Seasonal and spatial patterns of fungal populations. New Phytol. 138:131– 142. Morton, J.B., Benny, G.L. (1990) Revised classification of arbuscular mycorrhizal fungi (Zygomycetes)— a new order, Glomales, 2 new suborders, Glomineae and Gigasporineae, and 2 new families, Acaulosporaceae and Gigasporaceae, with an emendation of Glomaceae. Mycotaxon 37:471– 491. Morton, J.B., Redecker, D. (2001) Two new families of Glomales, Archaeosporaceae and Paraglomaceae, with two new genera Archaeospora and Paraglomus, based on concordant molecular and morphological characters. Mycologia 93:181– 195. Morton, J.B., Bentivenga, S.P., Bever, J.D. (1995) Discovery, measurement, and interpretation of diversity in arbuscular endomycorrhizal fungi (Glomales, Zygomycetes). Can. J. Bot.-Reυ Can. Bot. 73:S25– . S32. Muyzer, G., Dewaal, E.C., Uitterlinden, A.G. (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction amplifled genes coding for 16S ribosomal RNA. Appl. Enυ iron. Microbiol. 59:695– 700. Newsham, K.K., Fitter, A.H., Watkinson, A.R. (1995) Multi-functionality and biodiversity in arbuscular mycorrhizas. Trends Ecol. Eυ 10:407– ol. 411. O’ Connor, P.J., Smith, S.E., Smith, E.A. (2002) Arbuscular mycorrhizas influence plant diversity and community structure in a semiarid herbland. New Phytol. 154:209– 218. O’ Donnell, K., Lutzoni, F.M., Ward, T.J., Benny, G.L. (2001) Evolutionary relationships among mucoralean fungi (Zygomycota): Evidence for family polyphyly on a large scale. Mycologia 93: 286– 297. Orita, M., Iwahana, H., Kanazawa, H., Hayashi, K., Sekiya, T. (1989) Detection of polymorphisms of human DNA by gel electrophoresis as single strand conformation polymorphisms. Proc. Natl Acad. Sci. USA 86:2766– 2770. Pringle, A., Moncalvo, J.M., Vilgalys, R. (2000) High levels of variation in ribosomal DNA sequences within and among spores of a natural population of the arbuscular mycorrhizal fungus Acaulospora colossica. Mycologia 92:259– 268. Redecker, D. (2000) Specific PCR primers to identify arbuscular mycorrhizal fungi within colonized roots. Mycorrhiza 10:73– 80. Rodriguez, A., Dougall, T., Dodd, J.C., Clapp, J.P. (2001) The large subunit ribosomal RNA genes of Entrophospora infrequens comprise sequences related to two different glomalean families. New Phytol. 152:159– 167.
Sanders, I.R. (2002) Ecology and evolution of multigenomic arbuscular mycorrhizal fungi. Am. Natural 160:S128– S141. Sanders, I.R., Fitter, A.H. (1992) Evidence for differential responses between host fungus combinations of vesicular arbuscular mycorrhizas from a grassland. Mycol. Res. 96:415– 419. Sanders, I.R., Alt, M., Groppe, K., Boller, T., Wiemken, A. (1995) Identification of ribosomal DNA polymorphisms among and within spores of the Glomales— application to studies on the genetic diversity of arbuscular mycorrhizal fungal communities. New Phytol. 130:419– 427. Schü ler, A., Gehrig, H., Schwarzott, D., Walker, C. (2001a) Analysis of partial Glomales SSU rRNA ? gene sequences: implications for primer design and phylogeny. Mycol. Res. 105:5– 15. Schü ler, A., Schwarzott, D., Walker, C. (2001b) A new fungal phylum, the Glomeromycota: phylogeny ? and evolution. Mycol. Res. 105:1413– 1421. Schwarzott, D., Walker, C., Schü ler, A. (2001) Glomus, the largest genus of the arbuscular mycorrhizal ? fungi (Glomales), is nonmonophyletic. Mol. Phylogenet. Eυ 21:190– ol. 197. Simon, L., Lalonde, M., Bruns, T.D. (1992) Specific amplification of 18S fungal ribosomal genes from vesicular-arbuscular endomycorrhizal fungi colonizing roots. Appl. Enυ iron. Microbiol. 58: 291– 295. Simon, L., Bousquet, J., Levesque, R.C., Lalonde, M. (1993a) Origin and diversification of endomycorrhizal fungi and coincidence with vascular land plants. Nature363:67– 69. Simon, L., Levesque, R.C., Lalonde, M. (1993b) Identification of endomycorrhizal fungi colonizing roots by fluorescent single-strand conformation polymorphism polymerase chain-reaction. Appl. Enυ iron. Microbiol. 59:4211– 4215. Smith, S.E., Read, D.J. (1997) Mycorrhizal Symbiosis, 2nd edn. San Diego: Academic Press. Streitwolf-Engel, R., Boller, T., Wiemken, A., Sanders, I.R. (1997) Clonal growth traits of two Prunella species are determined by co-occurring arbuscular mycorrhizal fungi from a calcareous grassland. J. Ecol. 85:181– 191. Taylor, T.N., Remy, W., Hass, H., Kerp, H. (1995) Fossil arbuscular mycorrhizae from the Early Devonian. Mycologia 87:560– 573. Tehler, A., Farris, J.S., Lipscomb, D.L., K? llersj? M. (2000) Phylogenetic analyses of the fungi based on , large rDNA data sets. Mycologia 92:459– 474. Tester, M., Smith, S.E., Smith, F.A. (1987) The phenomenon of nonmycorrhizal plants. Can. J. Bot.-Reυ . Can. Bot. 65:419– 431. Trappe, J.M. (1987) Phylogenetic and ecological aspects of mycotrophy in the angiosperms from an evolutionary standpoint. In: Safir, G.R. (ed) Ecophysiology of VA Mycorrhizal Plants, pp. 5– Boca 25. Raton, Florida: CRC Press. Turnau, K., Ryszka, P., Gianinazzi-Pearson, V., van Tuinen, D. (2001) Identification of arbuscular mycorrhizal fungi in soils and roots of plants colonizing zinc wastes in southern Poland. Mycorrhiza 10:169– 174. van der Heijden, M.G.A., Boller, T., Wiemken, A., Sanders, I.R. (1998a) Different arbuscular mycorrhizal fungal species are potential determinants of plant community structure. Ecology 79: 2082– 2091. van der Heijden, M.G.A., Klironomos, J.N., Ursic, M., Moutoglis, P., Streitwolf-Engel, R., Boller, T., Wiemken, A., Sanders, I.R. (1998b) Mycorrhizal fungal diversity determines plant biodiversity, ecosystem variability and productivity. Nature 396:69– 72.
van Tuinen, D., Jacquot, E., Zhao, B., Gollotte, A., Gianinazzi-Pearson, V. (1998) Characterization of root colonization profiles by a microcosm community of arbuscular mycorrhizal fungi using 25S rDNA-targeted nested PCR. Mol. Ecol. 7:879– 887. Vandenkoornhuyse, P., Husband, R., Daniell, T.J., Watson, I.J., Duck, J.M., Fitter, A.H., Young, J.P.W. (2002) Arbuscular mycorrhizal community composition associated with two plant species in a grassland ecosystem. Mol. Ecol. 11:1555– 1564. White, T.J., Bruns, T., Lee, S., Taylor, J. (1990) Amplification and direct sequencing of fungal ribosomal genes for phylogenies. In: innis, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (eds) PCR Protocols: a Guide to Methods and Applications, pp. 315– 322. New York: Academic Press.