Fine-scale diversity and distribution of ectomycorrhizal fungal mycelium in a Scots pine forest

Authors


Summary

  • Ectomycorrhizal (ECM) mycelium is a key component of the ectomycorrhizal symbiosis, yet we know little regarding the fine-scale diversity and distribution of mycelium in ECM fungal communities.
  • We collected four 20 × 20 × 2-cm3 (800-cm3) slices of Scots pine (Pinus sylvestris) forest soil and divided each into 100 2 × 2 × 2-cm3 (8-cm3) cubes. The presence of mycelium of ECM fungi was determined using an internal transcribed spacer (ITS) database terminal restriction fragment length polymorphism (T-RFLP) approach.
  • As expected, many more ECM fungi were detected as mycelium than as ectomycorrhizas in a cube or slice. More surprisingly, up to one-quarter of the 43 species previously detected as ectomycorrhizas over an area of 400 m2 could be detected in a single 8-cm3 cube, and up to three-quarters in a single 800-cm3 slice. ECM mycelium frequency decreased markedly with depth and there were distinct ‘hotspots’ of mycelium in the moss/F1 layer.
  • Our data demonstrate a high diversity of ECM mycelium in a small (8-cm3) volume of substrate, and indicate that the spatial scale at which ECM species are distributed as mycelium may be very different from the spatial scale at which they are distributed as tips.

Introduction

Below-ground mycelial systems of ectomycorrhizal (ECM) fungal communities are extensive and can comprise up to 30% of the total microbial biomass (Högberg & Högberg, 2002) and up to 80% of the ECM biomass (Wallander et al., 2001, 2010) in boreal forest soils. ECM mycelial systems are also known to be an essential component of nutrient capture and carbon turnover (Smith & Read, 2008).

Despite the fact that we know that ECM mycelium is important, we know relatively little about how it is distributed in soil, and how the spatial distribution of mycelium relates to the spatial distribution of colonized root tips. Koide et al. (2005) found that the species composition of an ECM community, determined by the analysis of root tips or mycelium, was not the same and, for those species which occurred as both tips and mycelium, the frequency of detection was different. A number of studies have demonstrated vertical niche differentiation of ECM mycelial communities in relation to soil horizons (Dickie et al., 2002; Landeweert et al., 2003; Lindahl et al., 2007). Dickie et al. (2002) investigated the vertical distribution of ECM mycelium in the forest floor of a Pinus resinosa plantation and showed that, although some ECM species were found in all layers of the forest floor, others were more associated with either litter or more decomposed material. Landeweert et al. (2003) observed a similar vertical distribution pattern of ECM mycelium in a Pinus sylvestris forest, whereas Lindahl et al. (2007) found the mycelium of ECM fungi at all depths in a P. sylvestris forest soil profile, although they were more frequent in more decomposed litter and humus than in the surface litter and moss.

Using a spatially explicit sampling design, Genney et al. (2006) confirmed that the frequency of individual ECM species as mycelium was not the same as their frequency as tips, but they also showed that the spatial distribution of mycelium of a given species did not correspond to the distribution pattern of tips. For example, a Cortinarius sp. was much more frequent as mycelium than as tips and, although mycorrhizas were found between depths of 2 and 8 cm, the majority of the mycelium was found between 0 and 4 cm. It is important to emphasize that Genney et al. (2006) investigated the frequency and distribution of the mycelium of those ECM species which were also present as ECM tips in the sampled soil volume. They did not report the incidence of mycelium of ECM species which they did not encounter as tips. However, information on the frequency and distribution of the mycelium of these species is essential to obtain an understanding of ECM fungal community assembly processes.

With this in mind, we return to the samples analysed by Genney et al. (2006), but, in this case, present data on the entire ECM mycelial community, not just those species which occurred as ectomycorrhizas in the samples. Our aim was to investigate the fine-scale diversity and distribution of mycelium in a P. sylvestris ECM community and to determine the following.

  • In the small (8-cm3) soil volumes sampled by Genney et al. (2006), does most of the ECM mycelium arise from those species present as tips or is there a spatial mismatch between the ECM species colonizing root tips and those present as mycelium?
  • To what extent does mycelium of different ECM species co-occur in the same 8-cm3 volume of soil, and is there evidence for spatial structure in the distribution of ECM mycelium at the 2–20-cm scale of this study?
  • How many ECM species can we detect as mycelium in the 8- and 800-cm3 samples collected in this study, and what proportions of the ECM species known to occur in the 400-m2 plot from which the samples were taken do they represent?

The samples analysed were those described by Genney et al. (2006). Briefly, four 20 × 20 × 2-cm3 vertical slices of soil were extracted from a 400-m2 research plot in a P. sylvestris plantation, and each divided into 100 2 × 2-cm2 cubes. Within each cube, we identified the ECM species present as mycelium using terminal restriction fragment length polymorphism (T-RFLP) analysis and comparison of the community T-RFLP data with a database containing diagnostic terminal restriction fragments (TRFs) for a total of 117 specimens of ECM fungi (Pickles et al., 2012). The community of ECM fungi present as ectomycorrhizas in the 400-m2 plot has been the focus of intensive study (Pickles et al., 2010, 2012). We were therefore able to look, across soil horizons, not only at the number of species of ECM fungi present as mycelium in small 8-cm3 samples, but also at the frequency and distribution of those species in relation to the ECM species known to form ectomycorrhizas across the whole 400-m2 plot.

Materials and Methods

Site description and soil sampling

Soil samples were collected from Culbin forest, which is located on the southern shores of the Moray Firth, north-east Scotland (57°38′08″N, 03°42′07″W). Within the forest, a uniform 20 × 20-m2 plot of 125-y-old Pinus sylvestris L. was selected, as described in Genney et al. (2006). The soil at this site has a weakly developed podzolic profile composed of organic horizons (bryophyte/litter (L; c. 0–2 cm), fermentation (F; c. 2–4 cm) and humic (H; c. 4–12 cm)) over an aeolian sand mineral horizon (Gauld, 1981). Within the 20 × 20-m2 plot, four vertical soil slices (20 × 20 × 2 cm3) were extracted and a digital image of the slice was captured for future reference. The soil slices (and the resulting 2 × 2 × 2-cm3 cubes (see later)) analysed here are the same as those used in Genney et al. (2006).

Each slice was divided into 2 × 2 × 2-cm3 cubes (yielding 100 soil cubes per slice), the roots were removed and the remaining soil from each cube was stored at −20°C before DNA extraction (see Genney et al., 2006). The 2-cm cubes were allocated to different substrate layers as follows: (1) moss/F1, living and dead moss incorporating unfragmented and unconsolidated pine needles, twigs and reproductive structures; (2) F1/F2, consolidated litter in various states of fragmentation and decay, mostly consisting of partially fragmented needles; (3) mineral soil, sand with varying degrees of colloidal organic matter staining. It should be noted that a humified ‘H’ layer, in which original plant structures are not identifiable, was not present at this site. Where cubes fell on the border between two layers, they were allocated to the layer contributing most to the cube (Fig. 1a).

Figure 1.

(a) Photographs of the four soil slices showing layer boundaries (dotted line; moss/F1, F1/F2, mineral). (b) Maps of ectomycorrhizal (ECM) species richness in each of the four soil slices. (c) ECM species accumulation curves plotted by soil layer for each soil slice. Moss/F1, closed circles; F1/F2, open circles; mineral, triangles.

Identification of ECM mycelium in soil cubes by T-RFLP

DNA was extracted from the root-free soil cubes using the method of Griffiths et al. (2000). Soil cubes from the organic horizons were first ground with liquid N2 to homogenize the sample. A subsample of 1 g was then incubated at 65°C for 45 min in 600 μl of 5% cetyltrimethylammonium bromide (CTAB) buffer. Cubes from the sand horizon were macerated for 2 × 30 s at 5.5 m s−1 using Lysing matrix-E tubes (Qbiogene, Cambridge, UK) in a FastPrep bead beating system (Bio-101, Vista, CA, USA). T-RFLP analyses were performed for each soil cube DNA extract as described in Genney et al. (2006). Briefly, DNA from each was amplified using the fluorescently labelled primers ITS1F (6-FAM) and ITS4 (HEX; White et al., 1990; Gardes & Bruns, 1993) in a 50 μl reaction mix that contained 1 μl DNA template, 1 × buffer (16 mM (NH4)2SO4, 67 mM Tris-HCl (pH 8.8 at 25°C), 0.01% Tween-20), 2.0 mM MgCl2, 250 μM deoxynucleoside triphosphates (dNTPs) (Bioline Ltd, London, UK), 20 pmol of each primer, 1 μl BSA and 2.5 U BIOTAQ polymerase (Bioline). The PCR conditions comprised an initial denaturation step at 95°C for 5 min, followed by 29 cycles of 95°C for 30 s, 55°C for 30 s and 72°C for 30 s, with a final extension step of 72°C for 5 min on a PTC-220 DYAD™ Thermal Cycler (MJ Research Inc., Waltham, MA, USA). PCR products were purified using the ChargeSwitch PCR clean-up kit (Invitrogen, Paisley, UK) before T-RFLP analysis with both HinfI and TaqI restriction endonucleases (Genney et al., 2006). TRF separation was performed on an ABI PRISM™ 3130xl genetic analyser (Applied Biosystems, Warrington, UK) using POP 4 and a 50-cm column with a 15 s injection time at 1.5 kV for 40 min at 60°C. A GS-500 ROX size standard (Applied Biosystems) was included in every sample run to facilitate accurate sizing of each detected TRF. TRF sizes for fragments > 50 bp, and with a peak height detection limit of > 50, were determined using GeneMapper V3.7 software (Applied Biosystems) and exported to TRAMP (Dickie et al., 2002) for comparison with reference TRF profiles in a T-RFLP database containing entries for 117 ECM specimens (Pickles et al., 2012), including ECM species previously detected as tips by Genney et al. (2006). Where > 50 TRFs were detected in a particular sample, the TRFs were ranked by peak height, and only the 50 highest were exported to TRAMP (Dickie et al., 2002). Positive identification of the presence of mycelium for species in the ECM T-RFLP database was confirmed if all available TRFs were detected within an error of ± 1.5 bp.

Data analyses

From the TRAMP output, a spreadsheet was constructed to show the presence/absence of species in each cube, and this was used to calculate the overall frequency of species, the frequency of species in each substrate layer and the number of species detected in each cube. The average number of species in a substrate layer as a function of the number of cubes sampled in that layer within a soil slice was computed using PC-ORD V5.0 (McCune & Mefford, 1999). The number of species in each cube was used to map the distribution of ECM species richness in the slices using Sigmaplot 11.0. To test whether the mycelium of individual ECM species was randomly distributed across the substrate layers, the observed frequency distribution of those species with an overall frequency of > 20 was compared with their expected distribution based on: (1) the distribution of 8-cm3 cubes across the layers and (2) the distribution of all ECM species, using χ2. Finally, the frequency of each species in each layer in each slice was computed, and the composition and structure of the ECM fungal community in the three substrate layers was compared by ordinating these frequency data using principal components analyses in PC-ORD.

Results

Successful amplification was achieved in 387 of the 400 possible 2-cm cubes (97%; Table 1). All the calculations below are based on cubes with successful amplification, except for the distribution maps of the number of ECM fungi in each cube (Fig. 1b), where the species richness of cubes in which amplification failed was set as the mean of the species richness in the touching cubes.

Table 1. Summary of the ectomycorrhizal (ECM) data generated for each soil slice
 Slice 1Slice 2Slice 3Slice 4Mean ± SE
  1. a

    As reported in Genney et al. (2006). Slice = 20 × 20 × 2 cm3; cube = 2 × 2 × 2 cm3.

Number of ECM fungi as tips in a slicea6756 
Number of ECM fungi as tips in a cubea4333 
Number of cubes with successful PCR amplification
Moss/F122212324 
F1/F230344314 
Mineral48403157 
Total (Nmax = 100)100959795 
Number of ECM fungi as mycelium17252924 
Number (%) of cubes with ECM fungi
Moss/F116 (73)18 (86)21 (91)17 (71)80 ± 5%
F1/F25 (17)31 (91)20 (47)1 (7)40 ± 19%
Mineral14 (29)20 (50)14 (45)11 (19)36 ± 7%
Total35 (35)70 (74)55 (57)29 (31) 
Mean number of ECM fungi per cube
Moss/F12.593.864.433.173.51 ± 0.40
F1/F20.332.440.770.140.92 ± 0.52
Mineral0.331.100.650.330.60 ± 0.18
Total0.832.191.601.02 
Maximum number of ECM fungi in a cube991110 
Frequency (F) of all ECM fungi
Moss/F1578110276 
F1/F21083332 
Mineral16442019 
Total8320815597 
%F of ECM fungi present as tips
Moss/F13530223330.0 ± 2.5
F1/F26059525055.3 ± 2.2
Mineral6952305351.0 ± 6.9
Total45463437 

We detected the DNA of 37 potential ECM species (Fig. 2, Table 1), and the number of species in each slice ranged from 17 to 29. Seven species corresponded to those recorded as ectomycorrhizas in these soil slices previously (Genney et al., 2006), whereas a further 21 species have been recorded in intensive morphotyping (Pickles et al., 2010) or T-RFLP typing (Pickles et al., 2012) studies of ectomycorrhizas in the surface organic horizons of the 20 × 20-m2 plot from which the four 20 × 20-cm2 slices were taken. The remaining nine species have not been recorded as ectomycorrhizas in the surface organic horizons at the site in any of these previous investigations (Fig. 2).

Figure 2.

Absolute frequency of ectomycorrhizal (ECM) fungal species as mycelium (number of 8-cm3 cubes in which species occurred) as detected by terminal restriction fragment length polymorphism (T-RFLP). Black bars, those species previously detected as root tips in the slices (Genney et al., 2006); dotted bars, those species previously detected as root tips elsewhere within the Culbin site (Pickles et al., 2010, 2012); white bars, those species never detected as root tips at the Culbin site.

In slices 1, 3 and 4, the greatest number of species was found in the moss/F1 layer (Fig. 1c), even when taking into account the differences in the number of cubes analysed in each layer. However, in slice 2, the species richness in the three layers was more similar.

The DNA of at least one ECM fungus was detected in 49% of the cubes. The percentage of cubes containing ECM fungi was significantly higher (80%) in the moss/F1 layer than in the F1/F2 (40%) or mineral soil (36%) layers (Table 1), although there were differences between the slices. Slice 2 (Fig. 1) had an overall higher percentage of cubes with ECM fungi (74%) than the other three slices (31–57%), and this was particularly marked in the F1/F2 layer (91% as opposed to 7–47%).

The maximum number of ECM fungi detected in a single cube across all four slices was 11 (range 9–11; Table 1). Overall, the mean number of ECM fungi in a cube was 1.40 ± 0.11, but the number was significantly higher in the moss/F1 layer (3.51) than in the F1/F2 (0.92) or mineral soil (0.60) layers (Table 1). Cubes containing higher numbers of ECM fungi were clustered (Fig. 1b) at a scale of a few centimetres, with cubes containing six or more ECM fungi largely restricted to the moss/F1 layer (except in the case of slice 2, where patches of high species richness were also present in the F1/F2 layer). Conversely, large volumes of the mineral soil were devoid of ECM fungi and this was not a result of PCR amplification (Table 1).

The overall frequency of detection of ECM fungi was markedly different between the slices, with more than twice as many ‘hits’ in slice 2 than in slices 1 and 4. In each slice, between 34% and 46% of the hits were from fungi which occurred as ectomycorrhizas in that slice (Genney et al., 2006). The percentage of hits from fungi which occurred as ectomycorrhizas was significantly lower (30%) in the moss/F1 layer than in the F1/F2 (55%) or mineral soil (51%) layers.

The rank order of species frequency was different in the three soil layers (Fig. 3) and the ordination of the species frequency indicated that the community of ECM fungi in the moss/F1 layer was different from that in the other two layers (Fig. 4). Examination of the frequency distribution of individual species (Fig. 5, Table 2) showed that two species, Tomentella bryophila and Cortinarius croceus, were more frequent in the moss/F1 layer than would be expected from the overall distribution of species, and that another three, Tricholoma flavovirens, Clavulinaceae sp. and Cadophora finlandia, were less frequent than would be expected.

Table 2. χ2-tests of whether the frequency distribution of a given species across the three soil layers differs significantly (shown in bold) from that expected based on: (1) the frequency distribution of cubes across those layers; and (2) the frequency distribution of all ectomycorrhizal (ECM) species across those layers
 (1) χ2 (P)(2) χ2 (P)
Distribution of cubes (= 387)  
Cubes with ECM mycelium (= 188) 26.558 (0.0001)  
All ECM fungi (F = 543) 383.950 (0.0001)
Tomentella bryophila (F = 28) 61.278 (0.0001) 9.263 (0.0097)
Cortinarius croceus (F = 20) 42.885 (0.0001) 5.953 (0.0510)
Tricholoma terreum (F = 20) 36.119 (0.0001) 3.940 (0.1394)
Cortinarius biformis (F = 50) 72.927 (0.0001) 5.221 (0.0735)
Amphinema cfr byssoides (F = 39) 35.434 (0.0001) 1.262 (0.5320)
Cenococcum geophilum (F = 33) 9.130 (0.0104) 5.126 (0.0771)
Amanita fulva (F = 40) 10.078 (0.0065) 4.050 (0.1320)
Tricholoma flavovirens (F = 23) 7.842 (0.0198) 28.706 (0.0001)
Clavulinaceae sp. (F = 45)2.633 (0.2680) 14.748 (0.0006)
Cadophora finlandia (F = 73) 21.393 (0.0001) 32.675 (0.0001)
Figure 3.

Combined rank order of relative frequencies of ectomycorrhizal (ECM) species as mycelium in the four soil slices plotted by soil layer: (a) moss/F1 layer; (b) F1/F2 layer; (c) mineral soil.

Figure 4.

Principal components analysis (PCA) ordination of ectomycorrhizal (ECM) species frequency in the moss/F1 (closed circles), F1/F2 (open circles) and mineral (open triangles) soil layers for each of the four soil slices.

Figure 5.

Frequency distribution of individual ectomycorrhizal (ECM) species in the moss/F1 (black bars), F1/F2 (dotted bars) and mineral (white bars) soil layers across all four soil slices. The χ2 statistics showing whether the frequency distribution of a particular species is significantly different from the expected frequency distribution based on the frequency distribution of cubes, or the frequency distribution of all ECM fungi, are given in Table 2.

Discussion

Here, we have shown that a given volume of a Scots pine forest soil contains many more ECM species as mycelium than are present as colonized root tips; the maximum number of ECM species detected as mycelium in an 800-cm3 soil slice was more than four times greater than that previously detected as tips (seven ECM species; Genney et al., 2006), and the maximum detected as mycelium in an 8-cm3 cube was nearly three times greater than that detected as tips (four ECM species; Genney et al., 2006). Although this difference between ECM tips and mycelium confirms previous observations (Koide et al., 2005), we have been able to show this here in a spatially explicit way. Although we accept that the nature of our sampling scheme (2-cm-thick vertical slices) means that we have no knowledge of which ECM species might be present as tips immediately adjacent to the samples analysed, we do not consider that the clear mismatch between tips and mycelium is an artefact of the sampling scheme. The marked clumping of ectomycorrhizas of many species (Pickles et al., 2010, 2012) gives some support to this view.

Subsequent to the sampling described here, an exhaustive sampling (441 soil cores; 5 cm in diameter × 5 cm in depth) of ECM root tips from the organic horizons of the 20 × 20-m2 plot from which our samples were taken detected a total of 43 species (Pickles et al., 2012). The maximum number of ECM species detected as mycelium in any single 800-cm3 soil slice was 29, and the maximum number in an 8-cm3 cube was 11. Therefore, approximately one-quarter of all ECM species detected colonizing tips in the plot can be detected in a single 8-cm3 cube of soil, and nearly three-quarters in a random 800-cm3 soil slice. These observations suggest that the spatial scales at which ECM species are distributed as mycelium are very different to the spatial scales at which they are distributed as tips.

We detected a total of 37 ECM species as mycelium across the four soil slices. Less than 50% of the ECM mycelium (based on frequency) was from the seven species previously detected as tips in the same slices (e.g. Cadophora finlandia, Cortinarius biformis, Clavulinaceae sp., Cenococcum geophilum; Genney et al., 2006). The relationship between mycelia and tip frequency distribution has been discussed previously by Genney et al. (2006). The proportion of mycelium derived from species forming tips was lowest in the moss/F1 layer. This may reflect the exploration strategies (Agerer, 2001) of the species occurring there, but we must also acknowledge that the potential for DNA from spores to affect the data would be highest in this layer. A further 41% of the mycelium came from species known to occur as tips in the organic layers of the 20 × 20-m2 plot (e.g. Amanita fulva, Amphinema cfr byssoides, Tricholoma flavovirens; Pickles et al., 2010, 2012). Surprisingly, 16% of the ECM mycelium belonged to nine species never detected as tips (Tomentella bryophila, Tricholoma terreum, Amanita muscaria, Tricholoma imbricatum, Amanita rubescens, Tomentella sp., Russula paludosa, Byssocorticium sp., Russula sp.; Fig. 2). There are several potential explanations for this observation. First, these species may be infrequent colonizers of root tips at the site and/or colonize roots just outside the main plot, and therefore would have remained undetected even with our previous extensive sampling at the site. Second, these species may have primarily been colonizing root tips in the mineral soil and therefore not detected by Pickles et al. (2012). ECM root tip communities have been shown to be vertically stratified (Rosling et al., 2003; Tedersoo et al., 2003), and spatial segregation between mycelium and tips of ECM species has been observed previously at this site (Genney et al., 2006). Third, some species may have remained undetected because of their morphological similarity to other ECM species at the site. For example, Tomentella bryophila forms black mycorrhizal root tips whose gross morphology is similar to that of Cenococcum (U. Koljalg, pers. comm.). Although both were detected as mycelium in this study, C. geophilum was the most abundant ECM fungus as tips in three previous studies at the site (Genney et al., 2006; Pickles et al., 2010, 2012) and, given the morphological similarities with other species, it is possible that this may have represented a species complex in these previous investigations. Finally, it is possible that some species (e.g. Amanita muscaria) may have been colonizing birch trees that were adjacent to the main 20 × 20-m2 plot. Although these represent plausible explanations of why we detected some ECM species as mycelium that have never been detected at the site as tips, we are also mindful of the technical limitations of T-RFLP (as discussed in Avis et al., 2006; Genney et al., 2006; Dickie & Fitzjohn, 2007), including the potential for T-RFLP to detect ECM species only present as spores (Dickie et al., 2002; Avis et al., 2006).

Some slices were more species rich than others. For example, slices 2 and 3 had the highest total number of ECM fungi as mycelium, the highest mean number of ECM fungi per cube (across all three soil layers and in total) and the highest frequency of ECM fungi (across all three layers and in total; Table 1). However, slices 2 and 3 also had the deepest F1/F2 layer, and so these observations may reflect the depth and/or decomposition status of the organic matter in each particular slice. There are also hotspots of ECM species richness at the scale of several centimetres, which are largely restricted to the moss/F1 layer (Fig. 1b). The exception to this is slice 2, where the hotspots are associated with decomposing wood, which may offer a more favourable physical/chemical environment for the growth of ECM mycelium. These hotspots indicate the presence of some spatial structure in ECM mycelial communities.

There was a general trend of vertical stratification in the mycelium of ECM fungi in the different soil layers. However, ordination analysis was only able to distinguish species composition in the moss/F1 layer from the F2 and mineral layers which clustered together in the ordination plot (Fig. 4). This appears to be a result of changes in ECM community structure rather than species richness, with a distinct subset of species (e.g. Tomentella bryophila, Cortinarius croceus, Tricholoma terreum, Cortinarius biformis and Amphinema cfr byssoides) appearing to dominate in the moss/F1 layer (Fig. 5). ECM frequency decreased dramatically with depth, with the mineral soil having the lowest ECM frequency across all four soil slices. The highest ECM frequency was encountered in the moss/F1 layer for all soil slices, except for slice 2, which had equal ECM frequency in the moss/F1 and F1/F2 layers and the highest ECM frequency overall (Table 1).

In an important study, Lindahl et al. (2007) described changes in fungal mycelia community composition along vertical profiles through a P. sylvestris forest soil. They found that saprotrophic fungi were primarily confined to recently shed litter, whereas mycorrhizal fungi dominated in the underlying, more decomposed litter and humus. Our investigation differed from that of Lindahl et al. (2007) in several respects. The sampling scheme and protocols for fungal identification were different, and we did not gather information on species richness and frequency of saprotrophs. Although both studies were in P. sylvestris forest, the sites were very different, and there was no ericaceous understorey at our site. Furthermore, our organic layers were formed on wind-blown sand with no humus (H) horizons, whereas the site of Lindahl et al. (2007) had a mature podsol with well-developed horizon differentiation. These differences mean that any comparison of the findings in the two studies must be made with caution. However, there are important similarities, as well as one clear and intriguing difference.

We found 37 ECM fungi, more than twice the number (16) found by Lindahl et al. (2007). However, in both studies, Cortinarius species were an important (25–50%) component of the ECM fungal community and were particularly associated with moss and recently fallen needles. In our study, we did not distinguish between moss and recently fallen needle layers, which, on our site, graded into fragmented litter. However, our moss/F1 layer would appear to be broadly comparable with Lindahl's first five categories (needles, Litter 1, Litter 2 (moss), Litter 2 (needles), Fragmented Litter). If we accept this comparison, then a proportionately similar decline in ECM fungal species richness with depth (moss and younger organic matter > older organic matter > mineral soil) is found in both studies. The marked difference between the studies lies in the frequency with which the mycelium of ECM fungal species was detected in the different layers. Lindahl et al. (2007) found greater frequency of ECM fungi in the older humus layers, and argued that saprotrophic fungi outcompete mycorrhizal fungi from the upper part of the forest floor. As the substrate becomes depleted in available energy, the saprotrophs become less competitive, and are replaced by mycorrhizal fungi. Although we do not have data on saprotroph frequency, it is clear that, in our study, ECM fungal mycelia were more, not less, frequent closer to the surface, and appeared to be growing actively among the moss and relatively recently fallen needles. The reasons for this difference are not clear, but one could speculate that it may be related to such factors as differences between the study sites in litter quality, atmospheric nitrogen input, microclimate or the presence/absence of ericaceous species. Clearly, more studies are required before any general conclusions can be reached about the vertical frequency distribution of ECM fungal mycelium in forest floors.

We know very little about the turnover of ECM root tips and the time scales involved with colonization and turnover; however, Koide et al. (2007) have shown that, although the root tips of some ECM species show little variation in frequency across seasons (in a 13-month sampling period), others have highest frequencies in spring or summer. In addition, ECM root tips at the Culbin site have recently been shown to be highly dynamic across years (Pickles et al., 2010). We know even less about the turnover of ECM mycelium, although Koide et al. (2007) were able to cluster P. resinosa ECM fungi into three groups that displayed different temporal patterns of frequency as mycelium across seasons in a 13-month sampling period. Although our data provide a first glimpse of the fine-scale spatial structure of mycelium in a Scots pine ECM community, they represent a single point in time. Given that there are likely to be significant fluctuations in the abundance of ECM mycelium across different seasons (Koide et al., 2007) and years, these should be the focus of future fine-scale spatial studies, together with trying to gain a mechanistic understanding of the underlying processes giving rise to the spatial patterns observed.

Acknowledgements

We thank Pamela Parkin, Leanne Reid and Allan Wilson for their technical assistance and the Forestry Commission for access to Culbin Forest. We thank Liz Holden for her help with sporocarp identification. This work was funded by the Natural Environment Research Council (grant: NER/A/S/2002/00861) and the Scottish Government (Rural and Environment Research and Analysis Directorate). I.C.A. acknowledges support from the Australian Research Council (LX0881973).

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