Isolation and phylogenetic identification of a dark-septate fungus associated with the alpine plant Ranunculus adoneus


  • Christopher W. Schadt,

    1. Department of Environmental, Population and Organismic Biology, University of Colorado at Boulder, Boulder, CO 80309–0334, USA;
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  • Renée B. Mullen,

    1. Department of Conservation Planning, The Nature Conservancy, 2404 Bank Drive, Suite 314, Boise, ID 83705, USA
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  • Steven K. Schmidt

    Corresponding author
    1. Department of Environmental, Population and Organismic Biology, University of Colorado at Boulder, Boulder, CO 80309–0334, USA;
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Author for correspondence: Steven K. Schmidt Tel: +303 492 6248 Fax: +303 492 8699


  •  Dark-septate endophytic (DSE) fungi are ubiquitous in the roots of Arctic and alpine plants, yet very little is known about their phylogenetic identities or effects on their host plants.
  •  Several such fungi were isolated from the alpine snowbed plant Ranunculus adoneus in the Front Range of Colorado, USA; one isolate was chosen for detailed study. The ability of this isolate to re-colonize plant roots in pot cultures was assessed, and phylogenetic analyses were performed using small-subunit (SSU), 5.8S and internal transcribed spacer (ITS) 2 ribosomal DNA sequences.
  •  This isolate had the ability to produce root endophytic structures in pot cultures similar to those reported from other sources and observed in R. adoneus roots. SSU phylogenetic analyses showed this isolate to be related to a clade within the Euascomycetes containing the Leotiales and Erysiphales. In addition, SSU and 5.8S–ITS2 sequences showed high phylogenetic similarity to a variety of isolates reported from other plants of diverse geographical origins. Although most of these isolates remain unidentified, one closely related isolate was the anamorphic taxon Phialophora gregata.
  •  The results suggest that this DSE isolate might belong to the fairly closely related group of plant endophytes that have varied effects on the plants that they inhabit.


Dark-septate endophytes (DSE) are fungi that are common in the roots of Arctic and alpine plants over broad host and geographical ranges (Read & Haselwandter, 1981; Currah & VanDyk, 1986; Trappe, 1988; Stoyke & Currah, 1991; Jumpponen & Trappe, 1998a; Mullen et al., 1998) and may comprise a significant portion of Arctic and alpine soil biomass (Bissett & Parkinson, 1979; Flanagan, 1981). Despite the numerous observations of these fungi in plant roots, very little is known about their ecology, identity or effects on the plants that they inhabit (Jumpponen & Trappe, 1998a). Although some researchers have dubbed these fungi ‘pseudo-mycorrhizal’ (Wang & Wilcox, 1985), DSE fungi do not usually appear to have any adverse effects on plant health or appear to be associated with necrotic tissues in plant roots (O’Dell et al., 1993; Stoyke & Currah, 1993). Glasshouse studies have given mixed results as to the effects of these isolates on plant growth and nutrition. DSE isolates have shown the ability to increase foliar phosphorus (Haselwandter & Read, 1982; Jumpponen & Trappe, 1998b) and nitrogen (Jumpponen & Trappe, 1998b) concentrations, but effects on overall plant biomass appear to be mixed or dependent upon plant/fungus species combination and/or original soil nutrient status (Stoyke & Currah, 1993; Jumpponen & Trappe, 1998b).

It is likely that part of the confusion surrounding the role of DSE fungi in nature is that these organisms probably have broad taxonomic affiliations. In order to characterize these DSE fungi better for comparison between studies and research groups, knowledge of the degree of relatedness between isolates is critical. While many isolates of DSE fungi have never been identified, some researchers have observed rare events of asexual sporulation in culture. These sporulating isolates are commonly identified as Phialocephala fortinii Wang and Wilcox (Wang & Wilcox, 1985; Stoyke & Currah, 1991; Stoyke et al., 1992; O’Dell et al., 1993; Ahlich & Sieber, 1996; Harney et al., 1997; Jumpponen & Trappe, 1998a; Jumpponen, 1999). Most isolates, however, have not been observed to sporulate in culture. Owing to the absence of morphological clues for sorting isolates, some researchers have used methods such as restriction enzyme digests (e.g. RFLPs) of the internal transcribed spacer (ITS) ribosomal DNA (rDNA) region to assess the degree of relatedness between sporulating and nonsporulating isolates within studies. These studies have often concluded that a majority of isolates may have similar RFLP patterns to P. fortinii (Stoyke et al., 1992; Harney et al., 1997). While RFLP methods may be appropriate to compare strains in a particular study or laboratory, they do not lend themselves to comparisons between research groups with any certainty, because final confirmation of the identity of samples should involve running them side by side on agarose gels. Very few studies have characterized isolates with a DNA sequencing based approach. Sequence information not only allows opportunities for easy comparison of isolates between studies, but also allows researchers to assess the phylogenetic position of these organisms.

In the Colorado alpine, Ranunculus adoneus plants contain significant amounts of DSE fungi in overwintering roots as they are emerging during snowmelt. At this time soil temperatures are near freezing and new root growth has not yet occurred. This also corresponds with the time of maximum nitrogen uptake by R. adoneus and high fluxes of various forms of nitrogen into soil N pools (Mullen et al., 1998; Lipson et al., 1999). The objectives of the current study were to isolate and identify these abundant fungi from R. adoneus.

Materials and Methods

Collection, quantification and isolation

Ranunculus adoneus A. Gray plants were collected from Niwot Ridge Long Term Ecological Research (LTER) site in the Colorado Front Range (see the map in Brooks et al. (1996) and the photograph in West et al. (1999)). The study site is located at an elevation of 3510 m and was described by Mullen & Schmidt (1993). Roots were washed in de-ionized water and were separated into two subsamples, the first of which was placed in FAA (10 : 35 : 10 : 5 formalin, water, ethanol and acetic acid), and the second of which was surface-sterilized in 2.5% sodium hypochlorite solution and placed on Modified Melin–Norkrans (MMN) agar (Marx, 1969). After 1–2 wk, slow-growing, dark-pigmented fungi that emerged from the roots were isolated and transferred to fresh MMN agar plates. Isolates were obtained from two separate samplings: five isolates from 1993 and 17 isolates from 1994. Induction of the production of asexual spores by these isolates was attempted by cold storage at 4°C for periods of up to 2 yr, and additionally at 2°C and 7°C for up to 3 months. One isolate, DS16B from the 1994 sampling, was chosen for further investigation.

Root samples stored in FAA were stained using a modification of the method of Phillips & Hayman (1970) as described by Mullen & Schmidt (1993). Old roots that had overwintered and contained suberin and other constituents of mature cell walls were cleared in 10% potassium hydroxide (KOH) for at least 1 h. To ensure that roots were cleared, the KOH was replaced until the solution remained clear. Dark-septate fungi were then quantified as described by Mullen & Schmidt (1993).

Resynthesis with Zea mays

In order to verify that our cultured isolate was the same as those seen in plant roots, an experiment was carried out to test its ability to colonize sterile Zea mays roots in a growth chamber. Z. mays was used in this experiment instead of R. adoneus because attempts to germinate R. adoneus seeds were unsuccessful. Potting soil was autoclaved for 90 min on two consecutive days and was placed in pots which had been sterilized for 48 h in sodium hypochlorite solution (2.5%). In an experiment consisting of two control and two experimental pots, 10 4-mm plugs of isolate DS16B were placed in the rooting zone of the experimental pots. Z. mays seeds (IoChief, Rocky Mountain Seed Co., Denver, CO, USA), which had previously been surface-sterilized for 20 min in 30% hydrogen peroxide and germinated on 10% tryptic soy agar plates, were planted in inoculated or control pots (three plants each) and allowed to grow in a growth chamber (Conviron model E15, Controlled Environments Inc., Pembina, ND, USA) for 6–7 wk with a photon flux density of 500 µM m−2 s−1 at canopy level, a 15-h light period, and day and night temperatures of 20°C and 15°C, respectively. Plants were not given fertilizer or extra nutrients beyond those available after soil sterilization. Plants were harvested, and roots were washed and placed in FAA, stained as above, and shoots were dried and weighed. Fungi observed in the corn roots after staining were quantified using the method described above.

DNA extraction, PCR conditions and sequencing

Pure cultures of isolate DS16B were grown on ¼-strength malt extract agar (Difco, Detroit, MI, USA) for 3 wk. Fungal hyphae were scraped from the surface of four plates, placed in a 1.5-ml centrifuge tube, immersed in liquid nitrogen and lyophilized overnight. Approximately 50 mg of the dried tissue was then used for DNA extractions after the methods of Lee & Taylor (1990), with the addition of a mechanical disruption of cells by three cycles of freezing and grinding in liquid nitrogen. The sample pellet was allowed to air-dry overnight and then resuspended in 100 µl of TE (100 mM Tris, 100 mM EDTA, pH8.3-Sigma-Aldrich Inc., St. Louis, MO, USA) buffer.

A nested PCR approach was used to amplify the rDNA of DS16B using NS1 and ITS4 as external primers (producing an expected product of ~2400 bp) and NS17–NS22, NS3–NS8 and ITS1–ITS4 as internal primer sets (White et al., 1990; Gargas & Taylor, 1992). This approach ensured that the initial amplifications would not be inhibited by the presence of introns commonly associated with the 3′ end of the small-subunit (SSU) rDNA region of many Euascomycetes. Since ribosomal RNA may inhibit the successful PCR amplification of rDNA (Pikaart & Villeponteau, 1993), we pre-treated samples with RNAse ONE™ (Promega, Madison, WI, USA) prior to the first round of PCR amplification by incubating 50 µl of extract with 10 µl of 10X PCR buffer (Expand™ Long Polymerase System Buffer 3, Boehringer-Mannheim, Indianapolis, IN, USA) and 5 U of the RNAase for 30 min at 37°C in 0.6-ml thin-wall tubes. A master mix was then added to the same tubes so that the final concentrations were 500 µM of each deoxynucleotide triphosphate (dNTP), 0.5 µM of each primer, and 2.5 U of Expand™ Long Polymerase Mix (Boehringer-Mannheim) to a final volume of 100 µl with the buffer and template above. Thermocycle conditions for the first round of PCR consisted of an initial denaturation at 95°C for 2 min followed by 30 cycles of denaturation at 95°C for 1 min, annealing at 62°C for 1 min, and extension at 72°C for 2 min, as well as a final extension step of 72°C for 10 min.

Following this first round of amplification, the PCR product was gel-separated with 1.5% agarose and the band of approx. 2400 bp excised to exclude a smaller (approx. 750 bp) light, non-specific band. These gel slices were then purified using a QIAquick™ gel extraction kit (Qiagen, Valencia, CA, USA) and eluted in 100 µl of 20-mM Tris (Sigma) (pH 8.0). One microlitre of this purified DNA was then carried over to the second round of PCR amplifications. Each of the second round of 100-µl amplifications contained 200 µM of each dNTP, 0.5 µM of each primer (listed above) and 2.5 U of Pwo Polymerase, along with 10 µl of the supplied 10X PCR buffer with 20-mM magnesium sulphate (Boehringer-Mannheim). Thermocycle conditions for the second round of PCR amplifications consisted of an initial denaturation at 95°C for 2 min, followed by 30 cycles of denaturation at 95°C for 1 min, annealing at 55°C for 1 min, and extension at 72°C for 2 min, and a final extension step of 72°C for 10 min. Each of these amplification products was visualized on 1.5% agarose gels.

Two replicate 100-µl PCR products from each of the second round of amplifications were purified using the QIAquick PCR Purification Kit (Qiagen), eluted with 50 µl of water, combined and concentrated approximately to 20 µl using a DNA Speed Vac (Savant, Farmingdale, NY, USA). Two microlitres of each concentrated product were then used as template for sequencing reactions in combination with 3.2 pmol of each of the following primers: NS17, NS2, NS3, NS4, NS5 and NS22 (for NS17–NS22 product); NS3, NS4, NS5, NS22 and NS8 (for NS3–NS8 product); and ITS1, ITS2, ITS3 and ITS4 (for ITS1–ITS4 product), to a final volume of 5.2 µl. Each of these samples was sequenced using an ABI PRISM™ Dye-Termination Cycle Sequencing Kit and an ABI 377 automated sequencer (Applied Biosystems, Weiterstadt, Germany) at the Molecular Cellular and Developmental Biology–DNA Sequencing Facility at the University of Colorado, Boulder. The sequence was compiled and deposited in GenBank with the accession number AF168167.

Phylogenetic analysis and sequence comparisons

The SSU rDNA sequence for our isolate was compiled and aligned, and the alignment manually edited with those of 44 taxa (see Fig. 2, later, for GenBank accession numbers) with the aid of Sequencher™ 3.0 (Gene Codes Co., Ann Arbor, MI, USA). These taxa were chosen to represent as broadly as possible the phylogenetic diversity within the Euascomycetes, yet still contain nearly complete SSU sequences.

Figure 2.

Neighbour-joining bootstrap tree of small-subunit (SSU) data showing the position of dark-septate endophytic (DSE) isolate DS16B within the Euascomycetes (92% of 1000 bootstrap replicates) and the clade containing the Erysiphales, Leotiales and Thelebolaceae (E) (78% of all bootstrap replicates). The analysis also clearly delineated other recognized lineages within the Euascomycetes with moderate to strong bootstrap support, including the operculate discomycetes (A) (82% of 1000 replicates) and the major subgroup of Loculoascomycetes (B) (64% of 1000 replicates), as well as the clade containing the Lecanorales, the Eurotiales and the remainder of the Loculoascomycetes (C, D and B) (70% of 1000 replicates). Each taxon name is preceded by its GenBank accession number. The scale indicates the average number of nucleotide substitutions per position.

Owing to the large number of taxa and characters, only a neighbour-joining analysis was performed using PAUP (beta pre-release: Swofford, 1999) (Sinauer Associates, Sunderland, MA, USA). The evolutionary distances were calculated using a Kimura two parameter model and among site variation was assumed to follow a gamma shape distribution with a shape parameter equal to 0.5. All sites with ambiguous or missing data (including gaps) were excluded from the analysis. Branch lengths of effectively zero were collapsed during searching. Saccharomyces cerevisiae (Hemiascomycetes), Taphrina communis (Archaeascomycetes) and Neolecta vitellina (a basal Euascomycete: Landvik, 1995) were used as outgroup taxa. Bootstrap values are based on 1000 replications and include all groups compatible with 50% majority-rule consensus.

ITS and 5.8S rDNA sequences were used to retrieve similar sequences from GenBank (see Table 1 for accession numbers) using BLAST (Altschul et al., 1990). These sequences were then aligned with DS16B sequences as above and the percentage similarity for the overlapping regions determined. A portion of the 5.8S rDNA gene as well as ITS2 was then used to perform a neighbour-joining analysis as above except that rates for variable sites were assumed to be equal. Blumeria graminis (AF011283), Sclerotinia sclerotiorum (Z73799), Monilinia laxa (Z73787) and Monilinia fructicola (Z73777) were used as outgroup taxa. The ITS1 region was not included in this phylogenetic analysis, as it could not be reliably aligned overall the taxa represented in the analysis.

Table 1.  Percentage similarity and source of 5.8S, ITS1 and ITS2 sequences obtained through GenBank with DS16B sequences
Isolate Source/ID/Reference15.8SITS1ITS2Accession
  1. 1 References: (1) Carter et al. (1999); (2) Chen et al. (1996); (3) An et al. (1993); (4) unpublished data from GenBank; (5) Saenz & Taylor (1999); (6) Chambers et al. (1999). *N.D. = sequence not determined or reported.

Oat roots/None/(1)100%96%97%AJ246140
Oat roots/None/(1)100%96%95%AJ246144
Oat roots/None/(1)100%95%95%AJ246142
Oat roots/None/(1)100%95%95%AJ246143
Oat roots/None/(1)100%95%94%AJ246141
Soybean stems/Phialophora gregata/(4)100%83%92%AF132804
Soybean stems/Phialophora gregata/(2)100%82%92%U66731
Soybean stems/Phialophora gregata/(2)100%81%92%U66727
Soybean stems/Phialophora gregata/(2)100%81%92%U66729
Fesctuca psuedostems/None/(3)100%N.D.*92%X62986
Fesctuca psuedostems/None/(3)100%N.D.*90%X62980
Fesctuca psuedostems/None/(3) 98%N.D.*91%X62979
Fesctuca psuedostems/None/(3) 98%N.D.*91%X62991
Salal roots/Hymenoscyphus ericae/(4) 98%71%82%AF149068
Salal roots/Hymenoscyphus ericae/(4) 98%70%82%AF149069
Salal roots/None/(4) 98%69%82%AF149085
Salal roots/None/(4) 98%69%82%AF149084
Salal roots/None/(4) 98%70%81%AF149082
Liverwort rhizoids/Hymenoscyphus ericae/(6) 98%69%81%AF069439
Salal roots/None/(4) 98%69%80%AF149083
Liverwort rhizoids/Hymenoscyphus ericae/(6) 98%65%80%AF069505
Pinus roots/Phialophora finlandia/(5) 98%70%79%AF011327
Liverwort rhizoids/Hymenoscyphus ericae/(6) 98%69%79%AF069440
?????/Hymenoscyphus ericae/(4) 97%N.D.*84%AF081436
?????/Scytalidium vaccinia/(4) 97%N.D.*83%AF081439
?????/Hymenoscyphus sp./(4) 97%N.D.*81%AF081435
?????/Hymenoscyphus sp./(4) 97%N.D.*81%AF081440
Pinus roots/Phialocephala fortinii/(5) 97%65%77%AF011326


Our isolates had colony morphologies and hyphal structures similar to those of dark-septate fungi isolated by other researchers (as reviewed by Jumpponen & Trappe, 1998a). Colonies were grey to dark brown, although aerial hyphae in the centre of the colonies were lighter than submerged hyphae along the colony edges. No sporulation was observed in any of our isolates from either year, despite cold storage treatment periods of up to 2 yr at 4°C or at 2 and 7°C for periods up to 3 months.

To verify that our fungal isolate was the same as those observed in the roots, we inoculated sterile corn plants with isolate DS16B. After six weeks of growth, inoculated corn roots reached an average of 12% infection by DSE, whereas no infection was observed in control plants. Internal hyphae and ‘microsclerotia’ were morphologically very similar to typical DSE observed and reviewed by Jumpponen & Trappe (1998a), as well as those observed in R. adoneus collected from Niwot Ridge (Fig. 1a,b). For comparison, DSE infection rates of wild R. adoneus were greatest early in spring and reached peaks of approximately 29% (Mullen et al., 1998).

Figure 1.

Dark septate fungal colonization of root cells (bars, 100 µm). (a) Ranunculusadoneus root, showing microsclerotia characteristic of dark-septate endophytic (DSE) fungi filling a root cortical cell. (b) A Zea mays root inoculated with DSE isolate DS16B showing similar microsclerotia formations to those observed in wild Ranunculusadoneus plants.

PCR amplification of the rDNA gene of this isolate only proved possible after treatment with an RNA digesting enzyme. Before using RNAase, amplification was not achieved even with further 100- and 1000-fold dilutions of the template from extraction. After RNAase treatment we were able successfully to amplify and sequence nearly the entire 18S–ITS1–5.8S–ITS2 gene region using this nested PCRapproach.

The final SSU alignment used for phylogenetic placement of the DS16B isolate included 1546 characters. The alignment is homologous to that of S. cerevisiae (GenBank accession number J01353: Rubtsov et al., 1980) base positions 60–1669 of SSU rDNA, but excludes a highly variable region from base positions 637–740 which could not be unambiguously aligned.

The neighbour-joining analysis of SSU rDNA placed our isolate within the Euascomycetes with strong bootstrap support (92% of replicates) and within a clade (E in Fig. 2) containing members of the Erysiphales and Leotiales as well as the Thelebolaceae with moderate bootstrap support (78% of replicates; Fig. 2). The closest SSU rDNA sequence to DS16B was from an unidentified fungus isolated from the roots of winter wheat in Japan (Euascomycetes sp. K89, GenBank accession number AB016175) to which DS16B has over 99% identity and clustered with it in 100% of bootstrap replicates. The analysis also clearly delineated other recognized lineages within the Euascomycetes with moderate to strong bootstrap support, including the operculate discomycetes (A) (82% of 1000 replicates) and the major subgroup of Loculoascomycetes (B) (64% of 1000 replicates), as well as the clade containing the Lecanorales, the Eurotiales, and the remainder of the Loculoascomycetes (C, D and B) (70% of 1000 replicates).

The 5.8S–ITS2 region used for the second phylogenetic analysis included 251 aligned characters, beginning at a position homologous to that of S. cerevisiae base number 64 in the 5.8S rDNA gene (Rubin, 1973; GenBank accession K01048). Insertion–deletion sites within the ITS1 gene made alignments of this region ambiguous and prohibited the use of the ITS1 region in overall phylogenetic analyses. However, pairwise comparisons of this region with closely related sequences are shown in Table 1. The analysis showed DS16B to be closely allied with a variety of plant isolates of diverse host and geographical origins (Fig. 3), with the closest identified taxa being the anamorphic Phialophora gregata.

Figure 3.

Neighbour-joining bootstrap tree of 5.8S–ITS2 data showing the position of dark-septate endophytic (DSE) isolate DS16B within a monophyletic group of various plant isolates (97% of bootstrap replicates) and including the basal Phialocephala fortinii (79% of bootstrap replicates). Each taxon name is preceded by its GenBank accession number. The scale indicates the average number of nucleotide substitutions per position.


Phylogenetic analysis of DS16B placed it within the Euascomycetes and a clade containing the Leotiales, the Erysiphales and the Thelebolaceae ((formally within the Pezizales but excluded by Landvik et al. (1998)) (Fig. 2). The close relationship between these groups of ascomycetes has been noted several times previously (Saenz et al., 1994; Momol et al., 1996; Landvik et al., 1998), and an up-to-date classification of this group is needed as there seem to be several instances of paraphyly within this clade.

The most closely related organism to DS16B for which a full SSU sequence was available for comparison was an unidentified DSE fungus isolated from winter wheat roots in Japan (Euascomyces sp. K89, GenBank accession number AB016175). This fungus appears to be nearly identical to DS16B in culture and root infection morphology as well as in the SSU rDNA sequence. The Japanese isolate shared over 99% identity across the 18S alignment, and, of 11 mismatches, five were due to undetermined bases in one sequence or the other. Such high levels of identity indicate at least a very close relationship and perhaps even a conspecific nature. It is also of interest that K89 is not a pathogen of wheat, but rather it apparently antagonistically reduces the incidence (e.g. infection rates) and effects (reduction in biomass) of take-all disease (Gaeumannomyces graminis var. tritici) in wheat (Narita & Suzuki, 1991; S. Yamanaka, pers. comm.). These results mirror reports of a dark, sterile root endophyte that reduced the incidence of take-all disease in wheat (Speakman & Kruger, 1984), and numerous reports of such effects involving Phialophora radicicola (see Deacon, 1976; Speakman & Lewis, 1978), as well as other nonpathogenic Gaeumannomyces taxa (see Ward & Akrofi, 1994). All of these taxa, which might be considered DSE fungi, were omitted from review within Jumpponen & Trappe (1998a).

The position of our isolate in the SSU analysis is also nearly identical to that noted by others for P. fortinii. Using both parsimony and neighbour-joining analysis, LoBuglio et al. (1996) concluded that P. fortinii was probably associated with the Leotiales/Erysiphales. Unfortunately, we were not able to include this sequence in our analysis as we wanted to make full use of the nearly complete SSU sequence data that we obtained for our isolate, and the sequence reported by LoBuglio et al. (1996) was considerably less complete (1100 bp).
Jumpponen & Trappe (1998a) also used SSU rDNA sequences in a preliminary analysis of DSE isolates from a number of sources; however, the small region of 584 bp chosen for sequence analysis did not allow for a resolved placement of these fungi within the Euascomycetes, and their reported bootstrap values reflect this. Their study does suggest that DSE fungi are probably polyphyletic within the Euascomycetes; however, bootstrap support for the root of their tree was also not strong.

A unique aspect of our study is that we were able to perform analyses of both SSU and ITS sequences to narrow down the placement of our isolate as much as possible. Analysis of the two ITS regions and 5.8S rDNA using pairwise sequence comparisons also suggests that DS16B may be associated with a group of plant endophytic fungi with a wide geographical and host range but a fairly close phylogenetic relationship (Table 1). The 5.8S rDNA is identical to that of several isolates collected from oat roots in the United Kingdom, as well as soybean (identified as P. gregata) and fescue plants in different locations in the United States. The oat root isolates also displayed very high similarity in the ITS1 (95–96%) and ITS2 (94–97%). No ITS or 5.8S sequences of the Japanese wheat isolate (K89) that we found to be most closely related in SSU sequence are available at this time. Although levels of sequence similarity alone cannot confirm the relative taxonomic rank of these isolates (i.e. same species, genus, etc.), this small level of variation in the ITS sequences is well within the range of that reported within other species of fungi (O’Donnell, 1992; Carbone & Kohn, 1993; Farmer & Sylvia, 1998). For example, Table 1 lists the sequences of six isolates of Hymenoscyphus ericae and one of the teleomorph of this taxon Scytalidium vaccinia (Egger & Sigler, 1993). These sequences display only 93.3% similarity (6.7% divergence) in the ITS1 sequence, and 91.8% similarity (8.2% divergence) within ITS2.

The neighbour-joining analysis of the 5.8S–ITS2 sequences (Fig. 3) shows that the oat, soybean and fescue isolates in Table 1, as well as P. fortinii and our DSE isolate, form a monophyletic group supported in 79% of all bootstrap replicates. P. fortinii appears as a deeply divergent taxon within this group, whereas DS16B clusters well within the various plant isolates collected from other studies and associated with soybean, oat and fescue plants (97% of replicates). Hymenoscyphus spp. isolates, Gaultheria shallon (Salal) mycorrhizal isolates and Phialophora finlandia (another DSE fungus) form another, less well-supported, monophyletic grouping in this 5.8S–ITS2 analysis. The position of our isolate as basal and distinct from P. fortinii and relatively distantly related to P. finlandia further supports the polyphyletic origins of DSE fungi.

Although much has been written about the almost universal presence of DSE fungi in plant roots, we were none the less surprised that our isolate so readily infected the roots of a rather distantly related agricultural crop such as Z. mays. This finding indicates that at least this DSE fungus may indeed have the potential for a very broad host range. The fungal structures produced by our isolate by the end of the growth period very closely resembled those found in R. adoneus and many other wild plants (Jumpponen & Trappe, 1998a) with dark, supraficial and intracellular, septate hyphae that occasionally formed ‘microsclerotia’ (inflated, thick-walled cells) within root cortical cells (Fig. 1).

DSE make up a large component of Arctic and alpine soil fungi and although it is not known whether all DSE play similar functional roles in these and other ecosystems, their sheer abundance makes the need for further study important. Our ecophysiological studies have shown that DS16B has the ability to grow and actively to accumulate a variety of nitrogen forms at low temperatures in pure cultures (Mullen, 1995), and that DSE fungi may play an important role in early season nitrogen uptake by R. adoneus (Mullen et al., 1998). In addition, our SSU analysis has shown that DS16B is also very closely related to a winter wheat isolate that antagonistically reduces incidence and effects of the pathogenic take-all disease. Taken together, these results indicate that this particular group of DSE fungi could have a strong beneficial role in these systems. However, as DSE fungi are probably of diverse phylogenetic origins, molecular characterizations of DSE isolates under examination in future studies will be increasingly important for the interpretation of results between studies.


Support for this work was provided by National Science Foundation grants IBN9514123 and IBN9817164 and the Niwot Ridge LTER project (NSF 92117760). The authors would like to thank D. A. Lipson, D. Nemergut and R. E. Ley, as well as the two anonymous reviewers for comments on the manuscript.