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Fine scale distribution of ectomycorrhizal fungi and roots across substrate layers including coarse woody debris in a mixed forest Blackwell Publishing Ltd.

Leho Tedersoo1, Urmas Kõljalg1,2, Nils Hallenberg3 and Karl-Henrik Larsson3 1

Institute of Botany and Ecology, University of Tartu, 40 Lai Str., 51005 Tartu, Estonia; 2Institute of Zoology and Botany, Estonian Agricultural University,

181 Riia Str. 51014 Tartu, Estonia; 3Botanical Institute, Göteborg University, PO Box 461, SE 40530 Göteborg, Sweden

Summary Author for correspondence: Leho Tedersoo Tel: +372 56 654986 Email: [email protected] Received: 16 January 2003 Accepted: 2 April 2003 doi: 10.1046/j.0028-646x.2003.00792.x

• Ectomycorrhizal (ECM) fungi are widespread plant root symbionts in boreal forests, but information is lacking on the fine scale distribution of roots and fungi in substrate patches of different quality, including coarse woody debris (CWD). • Wood and soil cores were taken systematically both horizontally and vertically through decayed logs and underlying soil layers in a mixed forest. Root tips were counted and ECM fungi identified by morphotyping and sequencing. • The abundance of root tips and ECM fungi was highly variable on a 5-cm scale. Most species were replaced on a 50-cm scale. Detrended correspondence analysis demonstrated strong preference of resupinate thelephoroid and athelioid fungi and Sebacinaceae for CWD, whereas ascomycetes and euagarics appeared more frequently in mineral soil. Clavulicium delectabile was determined to be an ECM fungus for the first time. • ECM fungi occupy different niches and show variable distribution patterns. CWD plays an important role as a habitat both for roots and ECM fungi. We suggest sampling larger soil cores and selecting random root tips in future studies. Sequencing is a powerful tool in ECM community studies. Key words: Clavulicium, decomposed wood, ectomycorrhizal fungal community, fine scale, roots, sequencing, soil horizons, spatial distribution. © New Phytologist (2003) 159: 153–165

Introduction Ectomycorrhizal (ECM) fungi are preponderant plant root symbionts in boreal coniferous and mixed forests (Allen et al., 1995; Dahlberg, 2001), which cover vast areas in the northern hemisphere. Most fine roots and ECM of trees are aggregated in the uppermost 20 cm of soil (Schenk & Jackson, 2002), where nutrient circulation is most rapid. Mycorrhizal fungi are responsible for nutrient uptake (Read & Perez-Moreno, 2003) and provide species-specific benefits to plants (Perry et al., 1989; van der Heijden et al., 2003). Coarse woody debris (CWD), which consists of dead wood > 2.5 cm diameter, plays an important role, especially in forest and stream ecosystems (Harmon et al., 1986). In unmanaged forests CWD generally comprises 5–15% of total plant

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biomass (Brown, 2002; Krankina et al., 2002). Fine roots with ECM root tips are abundant in strongly decayed CWD (Harvey et al., 1978, 1979; Christy et al., 1982; Vogt et al., 1995; Goodman & Trofymow, 1998). Due probably to stimulated ECM activities, Väre (1989) reported increased plant growth following CWD amendment in a nursery in northern Finland. Saprotrophic fungi can create qualitatively different habitat patches (Pyle & Brown, 1999), which enhance the development of highly diverse communities of insects, mosses and late-successional fungi, including ECM fungi (Nordén et al., 1999). The remnants of brown rotted wood are abundant in the uppermost soil layer, hence different decomposition pathways determine the structure and patchiness of the whole organic horizon. The approximate diameter of a soil nutrient

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patch ranges from 20 cm in a deciduous woodland (Farley & Fitter, 1999) to 2 m in a coniferous forest (Liski, 1995). Because more fine roots are produced in patches of high nutrient content, their number and biomass is very variable in space and time (Pregitzer et al., 1993; Vogt et al., 1995). Single decayed logs include several nutrient patches (Pyle & Brown, 1999), which may influence the amount of roots and species composition of ECM fungi. Morphotyping and PCR–RFLP have been widely used to determine ECM fungi on plant roots, but these methods are too imprecise to discriminate closely related species (Kårén et al., 1997; Glen et al., 2001). Though more expensive and technically demanding, sequencing appears to be a powerful alternative to RFLP. In addition to generating a great number of characters for more reliable mycobiont identification, sequences also enable phylogenetic analysis. This possibility might prove useful to detect additional unexpected mycobionts that would remain unidentified by other methods (Selosse et al., 2002; Vandenkoornhuyse et al., 2002; Nilsson et al., unpublished). Most ECM community studies include samples from the organic horizon only, excluding information about the distribution patterns of ECM symbionts along the depth gradient (but see Heinonsalo et al., 2001). Though Agerer et al. (2002) developed micromapping to determine spatial relationships between ECM fungi in nature, there is limited information about fine scale patterns in ECM communities ( Yamada & Katsuya, 2001; Zhou & Hogetsu, 2002). Knowledge of the fine scale distribution of ECM is highly valuable to elaborate sampling designs for other ECM community studies. In this study we aim to: seek fine scale patterns of roots and ECM fungi; compare the distribution of ECM fungi in different soil horizons; assess the role of CWD for ECM fungi; and to assess the efficiency of direct sequencing (without an RFLP step) of root tips in ECM community studies.

Materials and Methods Study site Sampling was carried out in a 60 × 25 m rectangular site at Järvselja, eastern Estonia (58°17′N, 27°19′E; Fig. 1a,b). The site lies 36 m above sea level. The mean annual rainfall is 620 mm and mean annual temperature is +4.5°C. The vegetation type is Oxali-piceetum (Paal, 1997). The site is dominated by ECM hosts: Picea abies (L.) Karst. and Tilia cordata Mill. with a few Betula pendula Roth and Populus tremula L. P. abies, T. cordata and Sorbus aucuparia L. dominate the undergrowth. The forest floor is covered by Oxalis acetosella L., Vaccinum myrtillus L., V. vitis-idaea L., Rubus saxatilis L. and Hylocomium splendens (Hedw.) Schimp. in B.S.G. The soil is highly heterogenous due to abandoned drainage, waterlogged conditions in spring and mozaic substrate patches. The dominant soil type is podzol with organic

Fig. 1 (a) Map of the Baltic Sea region. An asterisk marks Järvselja. (b) Map of the sampling site (60 × 24 m) at Järvselja, Estonia. Living trees > 20-cm girth at breast height are shown. Trees > 40-cm diameter at breast height are indicated by larger symbols. Sampled logs are designated by straight lines. Plots are indicated by arrows and encircled numbers. Bar, 5 m (c) Sampling layout in a plot. A sample consists of three adjacent 5 × 5 × 5 cm subsamples. Samples were taken from four substrate layers. The two uppermost horizons comprise highly decomposed logs (coarse woody debris, CWD). Horizontal distance between samples, 50 cm; vertical distance, 7–50 cm.

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horizon (O), 7 cm; humus horizon (A), 20 cm; a transitional horizon between humus and eluvial horizons (AE), 7 cm; eluvial horizon (E), 40–60 cm; and B-horizon, below 75– 95 cm depth. Both A- and E-horizons feature a sandy texture. This site was selected because it is inhabited by several ECM hosts: spruce, 59.9% (breast area); hardwoods: linden, 22.0%; birch, 7.9%; and aspen, 6.5%. It contains 61.9 m3 ha−1 CWD > 10 cm diameter, including several well-decomposed logs, mostly spruce and aspen in decay stage IV (measured according to Renvall (1995)). Sampling Sampling was performed in August, 2001. Within the sampling site three plots were selected, each with one well decomposed log (see Appendix 1 for log characters). At three places along each log, samples (5 × 5 × 15 cm) were taken from the upper part of the log, the lower part of the log, the middle of the A-horizon, and the upper part of the E-horizon directly below. Each sample consisted of three adjacent 5 × 5 × 5 cm subsamples (Fig. 1c) cut with a woodsaw and/or a sharp knife. The horizontal distance between samples in the same plot was 50 cm. In all, 3 × 3 × 4 × 3 = 108 subsamples were taken, comprising 13.5 dm3 of substrate and 60 294 live root tips. Sampling was designed to assess the distribution patterns of roots and ECM fungi in both 5 cm and 50 cm scales. Subsamples were moistened if necessary, sealed in plastic bags, and stored at +4°C for up to 8 weeks. Subsamples were carefully sieved under tap water with a plastic sieve (1 mm mesh diameter) to remove any adhering soil particles. The fine roots with debris were laid in Petri dishes with abundant tap water. Only living root tips were separated by tree species and ECM morphotypes, and counted under an Olympus CO11 binoculare. ECM roots were distinguished from nonECM roots by the appearance of a fungal mantle. The presence of a Hartig net was not used as an ECM colonization criterium, because the Hartig net is not always well-developed, especially in young root tips. Herb roots were light-coloured, unlignified, narrower and not bearing ECM. Spruce roots could be distinguished from hardwoods by taste, smell, thickness, morphology of cross section, and shape of epidermal cells. Roots of different hardwood species never co-occurred.

Table 1 List of reference sequences obtained from sporocarps of ectomycorrhizal fungi

The criteria used to morphotype ECM were colour and surface of a mantle (cystidia, loose hyphae, laticiferous hyphae, etc.), and presence and type of emanating hyphae and rhizomorphs (Agerer, 2001). In case of ambiguity, morphotypes were separated into subtypes. Some root tips or clusters of each morphotype (depending on total amount) were excised and stored in distilled water at +4°C to take photos or isolate into pure cultures. Some additional root tips were preserved in FEA solution (formaline, 90%; ethanol, 5%; acetic acid, 5%) at room temperature to determine mantle structure (Agerer, 1991) and to take additional photos. Digital photos of plan view (magnification ×6–50) and mantle surface view (magnification ×1000) were taken with a Carl Zeiss binoculare Stemi 2000-CS and Axioskop 2 light microscope, respectively. Photos were adjusted with AxioVision 3.0 software. Sporocarp surveys were performed at 3–10 d intervals from August to October, 2001 and from May to October, 2002. All macrofungal sporocarps occurring at the study site were determined and listed. Voucher specimens of some species were dried and stored in the herbarium of the Institute of Zoology and Botany, Estonian Agricultural University (TAA). A specimen of Clavulicium delectabile (H.S. Jacks.) Hjortstam is deposited in the herbarium of Botanical Institute, University of Göteborg (GB). Selected fruiting bodies were later subjected to DNA extraction and sequencing (as mentioned in the following section) to obtain reference data (Table 1). Species nomenclature was used according to Hansen & Knudsen (1992, 1997, 2000). Higher level fungal taxonomy was applied according to Binder & Hibbett (2002) and Larsson (2002) for basidiomycetes, and LoBuglio et al. (1996) and O’Donnell et al. (1997) for ascomycetes. DNA extraction After counting, only the most healthy-looking and cleanest single root tips of each morphotype were carefully freed of reminiscent soil particles and fungal rhizomorphs, and mounted into Eppendorf tubes with 600 µl 2% CTAB buffer. Two separate tubes were prepared for each morphotype in a subsample. DNA was extracted according to the protocol of Savolainen et al. (1995), including GeneCleanTM treatment. PCR reactions were performed using Ready To GoTM Beads (comprising 1.5 units Taq DNA Polymerase, 10 mM Tris-HCl,

Species

Herbarium code

EMBL accession no

Clavulicium delectabile (H.S. Jacks.) Hjortstam Inocybe fastigiata (Schaeff.) Quel. Inocybe nitidiuscula (Britz.) Sacc. Lactarius trivialis (Fr. Fr.) Fr. Lactarius uvidus (Fr. Fr.) Fr. Russula betularum Hora Russula vinosa Lindbl.

KHL11147 TAA185038 TAA185039 TAA185040 TAA185041 TAA185042 TAA185043

AJ534896 AJ534933 AJ534934 AJ534935 AJ934536 AJ934537 AJ534938


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50 mM KCl, 1.5 mM MgCl2, 200 µM each dNTP and stabilizers, including BSA (Amersham Pharmacia Biotech.). 0.6–1 µl 20 µM primers were added. For the internal transcribed spacer (ITS) region a forward primer ITS4 (5′tcctccgcttattgatatgc-3′), ITS4B (5′-caggagacttgtacacggtccag-3′) or LR21 (5′-acttcaagcgtttcccttt-3′) was used in combination with a reverse primer ITS1 (5′-tccgtaggtgaacctgcgg-3′) or ITS1F (5′-cttggtcatttagaggaagtaa-3′). If the ITS sequence could not be matched in sequence databases, a larger fragment of 28S rDNA was amplified using a forward primer LR7 (5′tactaccaccaagatct-3′) together with a reverse primer LR0R (5′-acccgctgaacttaagc-3′) or CTB6 (5′-gcatatcaataagcggagg-3′). 6–10 µl DNA extract was added and finally bulked to 25 µl with sterile distilled water. PCR reactions were performed using Genius (Techne Ltd). An initial 3 min at 95°C was followed by 35 cycles of 30 s at 95°C, 30 s at 55°C, and 1 min at 72°C (2 section increment time for each following cycle; final cycle, 10 min) for each primer pair. 3.5 µl PCR products were run on 1% agarose gel with 6 µl ethidium bromide for 30–70 min. If PCR products resulted in several bands using primer pair ITS1/ITS4, faster moving bands (smaller DNA fragment) were separated from the gel as slower fragments represented DNA of hardwoods. For other primers, both products were separated. Cut gel was melted in TBE solubilizer and NaI (2 : 1 : 9 v/v/v) at 55°C, and subjected to GeneClean™ treatment. DNA giving a weak signal in the gel was treated similarily. Purified templates were reamplified with either the same or inner set of primers. Single PCR products were purified with QIAGEN PCR Purification Kit (QIAquickTM) according to the manufacturer’s instructions. DNA content was measured optically at 320 nm in GeneQuant pro. Sequencing was performed with the Ceq™ 8000 Genetic Analysis System (Beckman Coulter, Inc.) using primers ITS1, ITS2 (5′-gctgcgttcttcatcgatgc-3′), ITS3 (5′-gcatcgatgaagaacgcagc3′), ITS4, and CTB6, LR3R (5′-gtcttgaaacacggacc-3′), LR5 (5′-tcctgagggaaacttcg-3′), LR7 for ITS region and 28S rDNA, respectively. Raw sequences were checked for possible machine errors and primer disquality with Beckman Coulter software, and converted to Sequencher 3.×. format. Differently primed sequences of the same PCR product were merged to a contig using Sequencher 3.1 software (GeneCodes Corp.). Consensus sequences were composed, manually editing ambiguous readings, and queried in sequence databases at the National Center for Biotechnology Information (NCBI), European Molecular Biology Laboratory (EMBL), etc. using blastn algorithm (Altschul et al., 1997). ECM fungal isolates were considered as identified at species level when sharing > 97% ITS region sequence identity with a reference sequence in public databases or unpublished sequences. Otherwise the taxonomic interpretation depended on percentage sequence identities compared to one or several taxa, relative abundance of related taxa in the databases, the presence of unambiguously alignable ITS regions, the conservation of ITS sequence among different taxa, and the best 28S rDNA match(es) (see

Discussion). Eighty-five sequences, which represented 47 species of fungi, were derived from ECM root tips. The representative ITS and /or 28S rDNA sequences of each species were submitted to EMBL (accession numbers are indicated in Fig. 2). Data analysis To investigate the effect of depth gradient on fungal community composition, detrended correspondence analysis (DCA) was performed because it is based on simultaneous reciprocal averaging of both species and sample data. The analysis was performed with PC-ORD 4.01 (McCune & Mefford, 1999). Species abundance was converted to presence/absence due to the high number of rare species. Species data were merged at the level of major clades of basidiomycetes or orders of ascomycetes (see legend for Fig. 4), because we expected monophyletic groups of ECM fungi to possess similar ecological behaviour or properties (Chen et al., 2001, 2003; Webb et al., 2002). Three horizontally adjacent samples were merged into a sample triplet due to the high variation in occurrence of individual species. Frequence of clades/orders in sample triplets was applied in the analysis. Species of unknown affinity and sample triplets containing missing data were excluded. Regression analyses between root tip number, ECM colonization proportion and fungal parameters were performed to find significant correlations.

Results ECM fungal community There was a great variation in the number of root tips, relative abundance of fungal species and ECM colonization proportion in adjacent subsamples (Fig. 3). Cenococcum geophilum Fr. showed the most variable colonization pattern. The mean species richness was 3.65 ± 1.7 (mean ± ) and 4.68 ± 2.23 in a subsample and sample, respectively. Therefore, triplicating the subsample size increased the species richness by about one. When three random samples in a soil column or horizon in the same plot were pooled, 9.22 ± 2.99 and 10.79 ± 3.26 species were found, respectively. The mean species richness in a plot was 19.67 ± 3.79, which demonstrates a very limited overlap in the distribution of individual species across the 20–30 m between plots. Sample triplets taken from mineral and organic soil layers were readily distinguished by DCA according to major clades of basidiomycetes and orders of ascomycetes (Fig. 4). Sebacinaceae, athelioid and thelephoroid species acted as strong indicators of the organic substrate (CWD), whereas Pezizales, Helotiales and euagarics preferred mineral soil. Morphotyping and sequencing of ECM resulted in the recognition of 47 species of fungi (Fig. 5). Basidiomycetes and ascomycetes comprised 34 and 11 species, respectively,


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Fig. 2 Relative abundance of species of ectomycorrhizal fungi (ECM) in four substrate layers: (a) Upper coarse woody debris (CWD), (b) lower CWD, (c) humus (A) horizon, (d) eluvial (E) horizon. Counterclockwise from top: ascomycetes; clockwise: resupinate fungi, followed by other basidiomycetes. Different colours and lines represent species. Their EMBL accession numbers are indicated.


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Fig. 3 Relative abundance of root tips (bars), ectomycorrhizal fungi and nonectomycorrhizal root tips on a fine scale across four substrate layers. Vertical lines distinguish different plots. Samples without roots are not indicated. See Fig. 2 for legend and Fig. 1c for exact sampling layout. Note logarithmic scale for abundance of root tips only.

whereas two morphotypes remained unidentified due to recurrent failure in PCR. C. geophilum was ubiquitous, often being the only ECM mycobiont. C. geophilum colonized only single root tips without forming clusters. The other ECM fungi were dominant only in local patches and were generally restricted to one or two substrate layers or plots. Piloderma Fig. 4 Detrended correspondence analysis based on reciprocal averaging of data of sample triplets (triangles, see text for details) and frequency of major clades of ectomycorrhizal fungi (crosses). Numbered triangles indicate plots. Different horizons are shown in the legend. The effect of horizon as an environmental factor is represented by an arrow. Eigen values of DCA axes 1 and 2 = 0.242 and 0.114, respectively. CWD, coarse woody debris; Cg, Cenococcum geophilum. Corresponding genera of major clades/ orders are shown in brackets: basidiomycetes: athelioid (Amphinema, Piloderma, Tylospora), cantharelloid (Clavulicium, Clavulina), euagaric (Cortinarius, Hebeloma, Inocybe, Laccaria, unknown genus), russuloid (Lactarius, Russula), Sebacinaceae (Sebacina), thelephoroid (Tomentella); ascomycetes; Cenococcum geophilum, Eurotiales (Elaphomyces), Helotiales (Phialophora), Pezizales (Tuber, unknown genera).


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spp., Tylospora spp., Tomentella spp. and Sebacina spp. were found predominantly in CWD (Fig. 2). Except for C. geophilum, ascomycetes and euagarics were generally dominant in mineral horizons. Phialophora finlandia Wang & Wilcox was detected both as a contaminant endophyte and forming true ECM (cf. types f and g in Vrålstad et al. (2002)) on roots of deciduous trees only, mostly in the A-horizon. Tomentella (six spp.), Sebacina (four spp.), Inocybe (four spp.), Russula (three spp.) and Cortinarius (three spp.) were the most species-rich genera among ECM-forming fungi. Inocybe and Cortinarius spp. occurred in low numbers in one or two adjacent subsamples in the mineral soil. In addition to C. geophilum, only Tomentella sp1 occurred in all substrate layers, but the latter never exceeded 2% in abundance in any sample. This is the first report of ECM formed by Clavulicium delectabile. This species produces resupinate basidiocarps in debris and is closely related to ramarioid-clavarioid genus Clavulina (K-H Larsson, unpublished). Hosted by both spruce and birch, C. delectabile was found only in the upper layer of a white rotted aspen trunk. ECM of C. delectabile (Fig. 5a) is similar to those of other clavulinoid ECM with a strongly ramified and completely smooth appearance, type A plectenchymatous upper mantle layer and type L/M stretched pseudoparenchymatous inner mantle layer (cf. Agerer, 1991). Neither emanating hyphae nor clamp connections were observed. Beige ECM of C. delectabile was morphologically indistinguishable from ECM of Clavulina sp. (Fig. 5c). However, both species could be easily distinguished from ECM of Clavulina cf. coralloides, which possessed violaceous mantle colour (Fig. 5b). The number of ECM fungal species was strongly dependent on the abundance of root tips in a sample (Fig. 6). All spruce roots were visually colonized by ectomycorrhizal fungi. In deciduous trees we detected no trend in proportion of ECM colonization across substrate layers and between linden, aspen and birch. ECM colonization proportion did not correlate with any other parameters. Sporocarps of 46 species of ECM fungi were found at the site (Appendix 2). Russula, Cortinarius and Lactarius were the most abundant genera of ECM fungi. Only a few species were found in 2002, due a severe drought. Almost all species were found only once or twice in a restricted area. Only Lactarius deterrimus Gröger was found both above and below ground.

Discussion ECM community structure The relative abundance of ECM fungal species was highly variable on a 5-cm scale and nearly a complete change in species composition occurred both horizontally (50 cm) and vertically (except C. geophilum). We demonstrate that ECM communities may vary greatly on a fine scale inasmuch as species richness increased by one, when subsamples were merged. It is possible that a temporal change occurs among


ECM fungal species because we could find abundant dead ECM of some species (e.g. Tomentella stuposa), but all the living root tips were colonized by other species. Temporal changes in ECM communities occur during natural succession (Danielson, 1991; Baar et al., 1999), but seasonal and yearly fluctuations in stable communities remain poorly understood. The preference of major monophyletic groups of ECM fungi for different substrates was demonstrated by DCA (Fig. 4), although there were outliers and integral species in each group. Nevertheless, pooling the species data provided some generalizations and preliminary views on the potential niches of independently evolved monophyletic groups of ECM fungi. Differences in substrate preference might derive from different enzymatic arming to break down various substances, spreading strategies or functionality (Bruns, 1995; Agerer, 2001; Klironomos & Hart, 2001). There was no difference in the distribution of rhizomorph forming and smooth ECM types between mineral and organic substrate. However, our limited sampling effort does not allow drawing definitive conclusions and further evidence is required regarding functional diversity of major monophyletic groups of ECM fungi. The proportion of nonECM hardwood roots varied greatly and did not correlate with any other factors. Therefore, the proportion of ECM root tips might indicate the age of a particular root cohort (Black et al., 1998). Young nonECM roots might be colonized later inasmuch as it takes some weeks for ECM fungi to completely colonize a short root. Competition between arbuscular mycorrhizal and ECM fungi and dual colonization has been reported especially on roots of deciduous trees (van der Heijden, 2001). About 60% of the fungal species detected in ECM form no conspicuous sporocarps. These species colonized approx. 85% of total root tips. This pattern reflects either the true situation in a mixed type of forest, including CWD in the assay, or is derived from the identification of mycobionts by sequencing. Reference data of both ascomycetes and resupinate sporocarp forming basidiomycetes are very poorly represented in the RFLP databases (Kårén et al., 1997). This situation has probably strongly biased the ECM community composition towards mushroom-forming basidiomycetes in pine and spruce forests. Unfortunately, no comparative ECM fungal community studies of mixed forests have been carried out. However, the relative abundance of inconspicuous sporocarp forming fungi has been noted in several ECM community studies of monospecific coniferous forests (Flynn et al., 1998; Kõljalg et al., 2000). Thus, it is important to include these fungi in studies comparing sporocarps and ECM (Kernaghan & Harper, 2001). Clavulinaceae, Sebacinaceae and Pezizales comprise species that form a variety of sporocarp types (O’Donnell et al., 1997; Weiss & Oberwinkler, 2001). This study confirmed the wide distribution of Sebacinaceae as ECM symbionts (Glen et al.,

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Fig. 6 Dependence of fungal species richness on the number of root tips in the sample (n = 30; r = 0.574; P < 0.001). Triangles represent samples taken from different soil horizons (CWD, coarse woody debris).

2002; Selosse et al., 2002). The resupinate sporocarp former Clavulicium delectabile was determined to be an ECM symbiont for the first time. Based on our results, and as suggested by Horton & Bruns (2001), larger soil cores and random root tip picking could be useful for comparative studies on ECM fungal communities. This approach would reduce sampling effort, because not all root tips in a core would have to be found and counted. In addition, loose root tips would not need to be categorized by host tree species. The strong substrate preference of several ECM fungi underscores the need to sample different soil horizons in diversity studies. The role of coarse woody debris We detected 25 species of ECM fungi in three logs, which is more than described in previous studies in which samples were taken randomly (Harvey et al., 1979; Christy et al., 1982; Goodman & Trofymow, 1998). Christy et al. (1982) and Harvey et al. (1979) reported strong monodominance of an unknown species, whereas Goodman & Trofymow (1998) found C. geophilum and possibly two Piloderma spp. codominating inside stumps and logs. The results of the latter study are in accordance with our data. Both P. fallax and Tylospora spp. possess manganese and lignin peroxidase genes (Chen et al., 2001), which suggest their contribution to

degradation of remnants of decayed wood and soil humic polymers to acquire extra carbon and other nutrients from polyaromatic complexes. The extent to which ECM fungi can colonize less decomposed heartwood of CWD via rhizomorphs and emanating hyphae is unknown. Resupinate fungi may merely use woody debris as a support for building a down-facing hymenium. Tomentellopsis submollis (Svrcek) Hjortstam, Piloderma fallax and Tomentella crinalis (Fr.) M.J. Larsen, however, are able to grow saprotrophically on sterilized rotted wood of medium decomposition stages, where naturally no roots have been observed (L. Tedersoo, unpublished observations). Therefore ECM fungi may colonize dead wood by emanating hyphae with no root penetration and compete with saprotrophs for nutrients. Lindahl et al. (1999) and Leake et al. (2001) reported labelled nutrient translocation from a saprotrophinoculated wood plug to a plant via ECM hyphal network. These studies suggest that ECM fungi do not need to produce costly ligninases to acquire nutrients from CWD. Rather than direct nutrient capture, ECM fungi may obtain nutrients via absorbtion of oligomolecules exogenously decomposed by saprotrophic fungi. We found similar species of ECM fungi colonizing upper and lower parts of the log, although fewer ECM fungal species occupied the upper CWD stratum. Consequently, most ECM fungi are likely to disperse to the upper layer of a log together with host roots. We might expect more ECM fungal species in the upper parts of CWD if spores occur as a significant source of inoculum. Thus, we suggest that CWD does not act as a spore germination substrate for most ECM fungi. Direct sequencing Sequencing without an RFLP step proved to be a powerful tool in this community study. We could distinguish unambiguously between species after blast searching and manual aligning. However, detection of fungal species using molecular methods is dependent on success in DNA amplification. Correct sample storage and DNA extraction is important (Johansson, 2002). We obtained no sequences of obvious soil microfungi. Two unknown morphotypes were probably too senescent to extract high quality DNA. Of possible ascomycetous contaminants, Phialophora finlandia, Elaphomyces spp. and representatives of Pezizales have been reported as frequent ECM formers (Danielson, 1991; Baar et al., 1999; Vrålstad et al., 2002). In this study P. finlandia occurred as a typical ECM fungus, simultanuously colonizing several root tips endophytically – independent of their

Fig. 5 Plan views of ectomycorrhizas: (a) Clavulicium delectabile on spruce in white rotted aspen log; (b) Clavulina cf. coralloides on birch in E-horizon; (c) Clavulina sp. on aspen in A-horizon; (d) Piloderma sp. on spruce in decayed aspen wood; (e) Russula aff. sphagnophila on linden in brown rotted birch wood; (f) Sebacina sp1 on linden in brown rotted birch wood; (g) Sebacina sp2 on birch in white rotted aspen wood; (h) Sebacina sp3 on aspen in A-horizon; (i) Sebacina sp4 on linden in A-horizon; (j) Tomentella lilacinogrisea on linden in brown rotted birch wood; (k) Tomentella stuposa on birch in white rotted aspen wood; (l) Tomentella terrestris on linden in brown rotted birch wood; (m) Pezizales sp1 on aspen in A-horizon; (n) Tuber aff. maculatum on birch in E-horizon; (o) Elaphomyces sp1 on spruce in A-horizon. Bars, 1 mm.


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mycorrhizal status as also noted by Carlsen (2002) and Vrålstad et al. (2002). It is possible that P. finlandia could continue to grow at +4°C and increase its abundance inside root tips. Of the primer pairs used in the PCR reaction, the widely applied ITS1F/ITS4B and ITS1/ITS4 performed poorly. The latter coamplified DNA of deciduous trees, which could be either detected on gel or on a sequence chromatogram as a double signal. In several other studies of angiosperm ECM, ITS1/ITS4 performed well (Sønstebø, 2002). In our PCR conditions ITS1F/ITS4B gave poor results because they neglected all ascomycetes, Sebacinaceae, most athelioids (except Piloderma spp.) and several euagarics. Primer pairs ITS1/LR21 and ITS1F/LR21, which are not widely used, could amplify DNA even from the most problematic morphotypes with no contamination of plant DNA. In addition, these two primer pairs amplified a long DNA fragment (850– 950 bp) comprising the whole ITS region and approx. 300 bp of 28S rDNA. Simultaneous amplification of both conserved and variable regions creates a possibility to infer the phylogenetic position of the fungus at a higher taxonomic level if the ITS region cannot be matched in sequence databases. Additional sequencing of 28S rDNA could improve the determination of some unknown species, at least at the family or order level. Sequences of 18S, ITS and 28S rDNA regions are already available for many basidiomycetes, while 18S rDNA sequences are predominantly applied to study the phylogeny of ascomycetes. In addition to ITS, other rDNA regions and some functional genes might be used in future to resolve the phylogeny and to identify ECM fungi (Kretzer & Bruns, 1999).

Acknowledgements This study was funded by grants to U. Kõljalg from the Estonian Science Foundation, Grant nos. 4083 and 5232 and to N. Hallenberg by FORMAS, Grant no. 23.9/2001-1063. We thank Dr F. Martin and three anonymous referees for helpful comments on this paper. We are indebted to V. Aldén, H. Nilsson and I. Saar for help in sequencing. Dr R. Kjøller is thanked for sharing his unpublished sequences to identify some taxa. We are grateful to Drs J. Liira and B. Lindahl for suggestions concerning statistical analyses. We thank Dr R. Szava-Kovats for language correction.

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Appendix Appendix 1 Characters of sampled logs Log no.

I

II

III

Tree sp.

P. tremula

P. tremula

B. pendula

Decay stage

IV

IV

V

Decay type

White and brown rot, patchy;

white rot (c. 70%)

Brown rot

hard inner sapwood

hard inner sapwood

Bark

Present against forest floor, soft, roots frequently penetrating

Present against forest floor, soft, defragmented, roots frequently penetrating

Fully covered, tough, roots occasionally penetrating

Log diameter in sampled area (cm)

17–18

45 –60

26 –30

Moss cover

90%

100%

80%

Vascular plants growing on the log

C. arundinacea O. acetosella P. abies (seedlings) R. saxatilis S. aucuparia (seedlings)

D. carthusiana M. bifolium O. acetosella R. idaeus R. saxatilis S. aucuparia (seedlings)

A. platanoides (seedlings) O. acetosella P. abies (seedlings) S. aucuparia (seedlings)

The nearest potential hosts (diameter breast height (cm), distance (m))

P. abies (16.6, 2) P. abies (25.5, 2) P. abies (39.2, 4) P. tremula (71.7, 2.5)

B. pendula (9.2, 4) P. abies (13.0, 1) P. abies (12.1, 1) P. abies (18.5, 2) P. abies (11.8, 3) P. abies (15.9, 3) P. abies (50.0, 5)

P. abies (31.2, 1) P. abies (7.6, 3) T. cordata (26.8, 1) T. cordata (25.2, 1)

Appendix 2 List of fungal species producing macroscopic sporocarps at the study site Distance from plot I (m) Amanita fulva (Schaeff.) Pers. Amanita porphyria (Alb. & Schw. Fr.) Mlady Armillaria gallica Marxmüller & Romagn.a Cantharellus tubaeformis (Bull. Fr.) Fr.c Clavariadelphus sachalinensis (Imai) Cornera Clitocybe clavipes (Pers. Fr.) Kumm.a Clitocybe gibba (Pers. Fr.) Kumm.a Clitopilus prunulus (Scop. Fr.) Kumm.b Collybia butyracea (Bull. Fr.) Kumm.a Collybia distorta (Fr.) Quel.a Collybia peronata (Bolt. Fr.) Kumm.a Cortinarius anomalus (Fr. Fr.) Fr. Cortinarius armillatus (Fr. Fr.) Fr. Cortinarius bivelus (Fr. Fr.) Fr. Cortinarius cf. alboviolaceus Cortinarius pholideus (Fr. Fr.) Fr. Cortinarius sp1 Cortinarius sp2 Cortinarius sp3 Cortinarius sp4 Cortinarius sp5 Craterellus cornucopioides (L. Fr.) Pers.c Dermocybe cinnamomea (L: Fr.) Fr. Entoloma nidorosum (Fr.) Quel. Gloeophyllum odoratum (Wulfen: Fr.) Imaz.a Hebeloma cf. hiemale Helvella macropus (Pers. Fr.) Karst.

Distance from plot II (m)

Distance from plot III (m)

Voucher specimen

5 5 2 0.6

TAA182660

0.5

TAA182661

2

1


Research Appendix 2 Continued Distance from plot I (m) Heterobasidion parviporum Niemelä & Korhonena Hygrophoropsis aurantiaca (Wulf. Fr.) Schroet.a Hypholoma lateritium (Schaeff. Fr.) Schroet.a Inocybe mixtilis Britz. Inocybe nitidiuscula (Britz.) Sacc. Inocybe sp1 Lactarius camphoratus (Bull. Fr.) Fr. Lactarius deterrimus Gröger Lactarius fuliginosus (Fr. Fr.) Fr. Lactarius helvus (Fr.) Fr. Lactarius lignyotus Fr. Lactarius necator (J.F.Gmel. Fr.) Pers. Lactarius theiogalus (Bull. Fr.) S.F. Gray Lactarius trivialis (Fr. Fr.) Fr. Lactarius uvidus (Fr. Fr.) Fr. Leccinum cf. scabrum Leccinum scabrum (Bull. Fr.) S.F. Gray Leccinum variicolor Watl. Leucopaxillus gentianeus (Quel.) Kotl.a Lycoperdon perlatum Pers. Pers.a Lycoperdon pyriforme Schaeff. Pers.a Megacollybia platyphylla (Pers. Fr.) Kotl. & Pouz.a Mycena pura (Pers. Fr.) Kumm.a Mycena sp1a Mycena sp2a Panellus stypticus (Bull. Fr.) Karst.a Pluteus atricapillus (Batch) Fayoda Pluteus leoninus (Schaeff. Fr.) Kumm.a Polyporus varius (Pers.) Fr.a Rozites caperatus (Pers. Fr.) Karst. Russula betularum Horac Russula decolorans (Fr.) Fr. Russula delica Fr. Russula emetica (Schaeff. Fr.) Pers. Russula claroflava Grove Russula fragilis (Pers. Fr.) Fr. Russula rhodopoda Zvara Russula sp1 Russula sp2 Russula sp3 Russula vinosa Lindbl. Tricholoma inamoenum (Fr. Fr.) Gill.

Distance from plot II (m)

Distance from plot III (m)

Voucher specimen

2

2

TAA185039

2 5

2 3

5 3 2 2

2 TAA185040 TAA185041

3

0.5 0.1

0.5 0

TAA182662

2

TAA185042

0.5

4

a

1

1

TAA185043

Known to be saprotrophic or parasitic; bsuspected to be ectomycorrhizal; cectomycorrhizal species occurring on decomposed logs at the site.


165