Patterns of nitrogen and carbon stable isotope ratios in macrofungi, plants and soils in two old-growth conifer forests

Authors

  • Steven A. Trudell,

    1. Division of Ecosystem Sciences, College of Forest Resources, Box 352100, University of Washington, Seattle, WA 98195-2100, USA;
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  • Paul T. Rygiewicz,

    1. Western Ecology Division, National Health and Environmental Effects Research Laboratory, US Environmental Protection Agency, 200 SW 35th Street, Corvallis, OR 97333, USA
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  • Robert L. Edmonds

    Corresponding author
    1. Division of Ecosystem Sciences, College of Forest Resources, Box 352100, University of Washington, Seattle, WA 98195-2100, USA;
      Author for correspondence: Robert L. Edmonds Tel: +1 206 685 0953 Fax: +1 206 543 3254 E-mail: bobe@u.washington.edu
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Author for correspondence: Robert L. Edmonds Tel: +1 206 685 0953 Fax: +1 206 543 3254 E-mail: bobe@u.washington.edu

Summary

  • • To further assess the usefulness of stable isotope ratios for understanding elemental cycling and fungal ecology, we measured δ15N and δ13C in ectomycorrhizal and saprotrophic macrofungi, plants, woody debris and soils from two old-growth conifer forests in Olympic National Park, Washington, USA.
  • • Ecosystem isotope patterns were similar at the two forests, but differences existed that appear to reflect soil nitrogen availability and C allocation within the ectomycorrhizal symbioses. δ15N and δ13C of ectomycorrhizal and saprotrophic fungi differed in both forests, and a dual δ15N/δ13C plot provided the best means of distinguishing them. Within both groups, δ15N and δ13C differed among genera and species, and the difference in species composition was an important determinant of the different overall δ15N of the ectomycorrhizal fungi at the two forests.
  • • Variation in multiple ecophysiological traits such as organic N use, mycelial morphology and transfer of N to phytobionts appears to underlie the variation in the isotope signatures of ectomycorrhizal fungi.
  • • The varied isotope signatures of ectomycorrhizal fungi suggest considerable functional diversity among them. Life-history strategies could provide a framework for interpreting these patterns.

Introduction

Traditionally, the soil nitrogen cycle in forests has been thought to be governed by mineralization processes driven by bacteria. However, increasing evidence indicates that macrofungi, particularly those involved in mycorrhizal symbioses, play important roles in N and C cycling in temperate and boreal forests, and that the N cycle in these systems is more complex than previously thought (Lindahl et al., 2002; Read & Pérez-Moreno, 2003). For instance, it is now known that many plants can access organic forms of N, usually via their mycorrhizal symbionts (Näsholm et al., 1998, 2000; Aerts & Chapin, 2000). Ectomycorrhizal (EcM) fungi have been shown to utilize N directly from organic materials such as proteins (Abuzinadah & Read, 1986); leaf litter (Bending & Read, 1995; Pérez-Moreno & Read, 2000); pollen (Pérez-Moreno & Read, 2001a); collembolans (Klironomos & Hart, 2001); and nematodes (Pérez-Moreno & Read, 2001b), and to transfer acquired N to their plant partners. This allows ‘short-circuiting’ of the traditional N cycle, as the mineralization pathway need not be involved, and provides a means for close coupling of the N and C cycles.

Ectomycorrhizal fungi are taxonomically diverse, comprising at least 5000–6000 species. Most EcM trees are able to associate with many different mycobionts, and most EcM fungi are able to associate with many different phytobionts (Molina et al., 1992). This leads to a great taxonomic diversity of EcM associations. However, the factors responsible for this diversity are little understood and it is uncertain whether important links exist between taxonomic and functional diversity. Often it has been assumed that the apparently low degree of host-specificity in EcM associations indicates a high degree of functional redundancy among the mycobionts. At the same time, it has been proposed that the taxonomic diversity of EcM fungi reflects great diversity in function (Cairney, 1999). That some, but not all, EcM fungi can obtain N directly from soil organic matter suggests the latter view. However, ‘establishing biodiversity–function relationships remains one of the most intractable challenges in ecological research’ (Leake, 2001), and many more data are needed before the importance of such relationships can be assessed fully.

Unfortunately, soil organisms such as fungi and the processes they mediate are difficult to study. Direct field observation entails disturbance and provides little control over the myriad of variables in the system. Laboratory experiments allow more control and clearer observation but, because of the necessary simplification, their relevance to nature often is not clear (Read, 2002). Thus, elucidating the functional roles of fungi will require a variety of creative approaches. One promising methodology is natural abundance stable isotope ratios. Most biologically important elements occur as two or more stable isotopes, with one being far more abundant than the other(s). Fractionation of the isotopes by biological and physical processes leads to concentration differences in substances of biological interest, and these differences can provide insights into fluxes among organisms; between organisms and their abiotic environment; and among compartments of the abiotic environment. An important advantage in using natural abundance stable isotope ratios for ecosystem studies is their ability to present a time-integrated picture of functional processes which often are difficult to measure directly (Robinson, 2001).

Over the past 10 yr, numerous surveys of N and C stable isotope ratios in sporocarps of macrofungi have been reported (Gebauer & Dietrich, 1993; Handley et al., 1996; Lilleskov et al., 1997, 2002; Taylor et al., 1997; Gebauer & Taylor, 1999; Hobbie et al., 1999a, 2001; Högberg et al., 1999b; Kohzu et al., 1999; Chapela et al., 2001; Henn & Chapela, 2001; Trudell et al., 2001, 2003; Griffith et al., 2002; Horwath et al., 2002; Taylor et al., 2003), most carried out in north temperate or boreal, often conifer-dominated, forests. These have suggested several patterns: (1) δ15N in EcM fungi is comparable to that in mineral soil and greater than that in plant foliage and saprotrophic fungi; (2) δ15N and δ13C in both EcM and saprotrophic fungi are greater than those in their bulk substrates; (3) δ15N and δ13C in soils increase with depth; (4) δ13C in EcM fungi is less than that in saprotrophic fungi, but greater than that in plant foliage. However, confidence in the generality of these patterns, and our ability to draw inferences about the sources and magnitude of variation in stable isotope ratios, are constrained because, with the exception of Taylor et al. (2003), each of these studies has at least one of the following limitations: (1) many sporocarp samples are not identified beyond family or genus; (2) sample sizes are small (one or two observations per species, with each observation representing a single sporocarp); (3) few or no analyses of saprotrophic fungi and/or associated ecosystem pools such as plants and soil are included; and (4) data from different forest types are pooled.

The recent study by Taylor et al. (2003) represents a major advance, by including a large number of analyses with all fungi identified to species, replicate analyses of most species, and inclusion of many saprotrophic fungi plus leaves from associated phytobionts. Three important findings from that study are: (1) isotope signatures of fungus sporocarps varied by family, genus and species; (2) isotope data were most informative at the species level; and (3) species composition of the EcM fungi was important for determining their aggregate isotope signatures. Earlier, Lilleskov et al. (2002) provided evidence that isotope signatures can reflect ecophysiological function – specifically, that EcM fungi that can utilize organic N exhibited higher δ15N than did species restricted to mineral N sources. Hobbie and colleagues (Hobbie et al., 1999a, 1999b, 2000, 2001; Hobbie & Colpaert, 2003) have presented a series of increasingly refined models relating δ15N in fungus sporocarps and plant foliage to soil N status and ecosystem processes. For instance, based on theoretical grounds, field measurements and a laboratory study, they propose that EcM fungus–plant differences in δ15N in part reflect the proportion of mycobiont N transferred to phytobiont(s) and that this, in turn, is influenced by allocation of C from the phytobiont(s) to the mycobiont. Other investigations have focused on the mechanisms that underlie the isotope effects (Högberg et al., 1999a; Kohzu et al., 2000; Emmerton et al., 2001; Henn & Chapela, 2004). Taken together, the results of these studies allow us to begin to understand the processes that produce the different isotope signatures of EcM and saprotrophic fungi, and their associated soil and plant pools. However, there remains a need for more extensive data on sporocarps and other pools from a greater variety of habitats to test the generality of the existing observations and conclusions.

To further assess the usefulness of stable isotopes for understanding elemental cycling and fungal ecology in forests, we have been studying the N and C stable isotope patterns in macrofungi, plants and soils in two old-growth conifer forests from climatically different areas of the Olympic Peninsula, western Washington, USA. Here we: (1) present the largest set of N and C stable isotope data on macrofungi and associated ecosystem pools reported to date; (2) provide additional support for the principal conclusions of Taylor et al. (2003) and extend their observations to additional taxa in a different forest setting; (3) relate the observed stable isotope patterns to differences in the species composition of the macrofungi and discuss possible ecophysiological bases for the relationship; and (4) compare our data with the model of Hobbie and Colpaert (2003) to assess its usefulness for interpreting field isotope measurements.

Materials and Methods

Study areas

The study was conducted in two areas within Olympic National Park, western Washington, USA. The first, lower Deer Park Road (DP), is located on the north-eastern side of the Olympic Peninsula, c. 15 km south of the Strait of Juan de Fuca (47°59′N, 123°19′W, 750–1025 m a.s.l.). The second, the Hoh River Valley (Hoh), is located on the western (windward) side of the Peninsula, c. 30 km east of the Pacific Ocean coast (47°50′N, 124°02′W, 170–250 m a.s.l.). The DP and Hoh areas are c. 55 km apart on opposite sides of the Olympic Mountains. Mean annual precipitation is c. 1000–1300 mm at DP and c. 3300–3600 mm at Hoh. Mean annual, January and July temperatures at DP are 9, 2 and 16°C, respectively; those at Hoh are 10, 4.5 and 16°C.

The dominant tree species at DP and Hoh are similar, with western hemlock (Tsuga heterophylla[Raf.] Sarg.) very abundant in both areas. Douglas fir (Pseudotsuga menziesii[Mirb.] Franco) and western redcedar (Thuja plicata Donn.) occur in both areas, but are more abundant (especially Douglas fir) at DP. Sitka spruce (Picea sitchensis[Bong.] Carr) is present only at Hoh, and is abundant there. Maximum ages of trees are > 300 yr at DP (O'Dell et al., 1999) and > 600 yr at Hoh (Edmonds et al., 1998). The understorey at DP consists primarily of mosses and scattered shrubs, especially salal (Gaultheria shallon Pursh, Ericaceae). Few herbaceous plants are present. The understorey at Hoh includes a greater variety of mosses, a higher density and diversity of shrubs, and a variety of herbaceous plants such as redwood sorrel (Oxalis oregana Nutt.), false lily-of-the-valley (Maianthemum dilatatum (Wood) Nels. & Macbr.), foam flower (Tiarella trifoliata L.) and western trillium (Trillium ovatum Pursh). Additional characteristics of the two areas have been described previously (Trudell et al., 2003; Trudell & Edmonds, 2004).

Sample collection and handling

The primary objective of sampling was to obtain five sporocarp collections per area of as many macrofungi as possible, plus samples from associated ecosystem pools, for stable isotope analysis. To capture as much of the variability in species occurrence as possible, sporocarps of macrofungi were collected at five sites in each of the two study areas. To standardize collecting effort, three 400 m2 plots were established at each site (1200 m2 total per site). Thus total sporocarp plot area was 0.6 ha at each of the two study areas. Plant, coarse woody debris (CWD) and soil samples were collected from areas immediately adjacent to the sporocarp plots (within c. 5–10 m) at the most productive site (in terms of sporocarp abundance) in each area. We visited the study areas 24 times from April 2000 to November 2001. Sporocarps were collected whenever present, soil samples five times, and plant and CWD samples once.

In this study, macrofungi are those producing epigeous sporocarps that are readily observable with the unaided eye: gilled mushrooms, boletes, gasteromycetes, polypores, corals, stipitate hydnums and large ascomycetes. No attempt was made to collect corticioid fungi. We did not make simultaneous multiple collections of a species within the same small area, thus collections made during a single sampling visit were probably from different mycelia. It is probable, however, that multiple collections were made from some mycelia during the course of the study.

Fresh leaves and twigs of Douglas fir (at DP), Sitka spruce (at Hoh), and western hemlock (at both areas) were cut with a pole pruner from the lower crown of individual canopy trees (five per species per area) in June 2001. Cores of sapwood and outermost heartwood from the same trees (one core per tree) were obtained with an increment borer. Needles were not separated by age class; they mostly represented years 1–3. Five samples each of CWD, representing two decay classes (2–3, moderately decayed and 4–5, highly decayed; Maser et al., 1979) were obtained per area in June 2001. The samples were made up of several subsamples scooped or chipped with a trowel from the bark-free surface of single downed conifer logs of undetermined species. Diameters of the logs ranged from c. 40–100 cm. Three soil samples per horizon, per area, were collected on five occasions (May, June, September, October, November 2001). Each sample was composited from three random subsamples taken with a trowel within c. 3 m radius of a random sample point. O-horizon samples encompassed the entire thickness of the horizon (Oi + Oe + Oa). Mineral samples comprised the upper 10 cm and, in the October sampling only, 10–20 cm samples were also collected. Sample handling has been described previously (Trudell et al., 2003; Trudell & Edmonds, 2004).

Names of plants follow Hitchcock and Cronquist (1973). Numerous literature sources were used for identifying fungi. Categorization of fungi by trophic types – EcM and soil (‘humus’, SapH), litter (SapL) and wood (SapW) saprotrophs – was based on information from Miller (1982), Singer (1986), and Molina et al. (1992).

Stable isotope analyses

Whole sporocarps from individual collections were analyzed. Whenever possible multiple sporocarps were included, thus the samples analyzed comprised from 1 to 104 sporocarps (mean = 5; median = 2). In cases where only portions of large sporocarps were used, they were sectioned longitudinally to preserve the relative proportions of stipe, hymenium and cap tissues. Stipes and whole caps from single sporocarps have been observed to differ in δ15N by c. 1–3‰ (Handley et al., 1996; Taylor et al., 1997). Analysis of dissected sporocarps of Leccinum scabrum, Paxillus involutus and Russula xerampelina during this study (data not shown) indicated that the cap–stipe difference was attributable mostly to enrichment of the hymenium tissue, not solely to the spores. Relative to cap and stipe trama, spores were enriched (c. 1.4‰) in R. xerampelina, but highly depleted (c.−5‰) in P. involutus. Sample preparation and analysis and quality assurance procedures have been described previously (Trudell et al., 2003; Trudell & Edmonds, 2004). Results of replicate analyses typically were within 0.2‰, both for δ15N and δ13C.

The stable isotope composition of the samples is expressed in differential notation as parts per thousand (‰) relative to a standard reference material. For example, for the two stable isotopes of N this is:

image(Eqn 1)

Atmospheric N and PeeDee Belemnite C were used as standards. Differences between the δ values of different pools are denoted by Δ, also in ‰.

Statistical analyses

Statistica 6.0 software (StatSoft, Oklahoma City, OK, USA) was used to calculate descriptive statistics (mean, SD) and perform statistical tests. Analytical variables were δ15N and δ13C. Separate comparisons were made for fungus trophic types, fungus genera within study areas, and fungus species within genera using one-way anova, followed by multiple comparison tests (Tukey's hsd for unequal n) at overall (experimental) α = 0.05. Comparisons between fungus trophic types (EcM-Sap) and study areas (DP-Hoh) were made using t-tests for independent groups. Additional comparisons between areas (DP-Hoh and Washington–Sweden) based on shared species were made using t-tests for dependent samples (paired t-tests). The degree of association between the isotope values among shared species of EcM fungi at the DP-Hoh areas and two sites studied by Taylor et al. (2003), Åheden and Stadsskogen, was examined using Pearson's product–moment correlation coefficient. The level of significance (α) for the t-tests and correlations was set at 0.05.

Because the numbers of macrofungus collections and species varied greatly among the sample plots and sites, all sporocarp data from the 15 plots at the five sites in each area were pooled, and all comparisons were made at the area (DP vs Hoh) level. Based on preliminary data from our study (Trudell et al., 2001), and data from Taylor et al. (2003) showing significant variation in δ15N among genera and species of EcM fungi, we used species means of fungi in comparisons involving trophic types, study areas and taxonomic groups. Thus in these cases n represents number of species. When making comparisons involving genera at study areas and species within genera, only genera with ≥ 3 species having ≥ 3 analyses at an area were included. Means for nonfungus ecosystem pools are based on individual sample values.

Results and Discussion

Stable isotope analyses

Altogether, we analyzed 731 sporocarp collections (391 from DP, 340 from Hoh) representing 152 species (89 at DP, 96 at Hoh, 32 at both) from 56 genera (37 at DP, 44 at Hoh, 25 at both). Mean δ15N and δ13C for each species, by area, are given in Appendix 1. Of the analyses/species means at DP, 327/65 were from EcM taxa, 11/3 SapH, 21/7 SapL, 31/13 SapW, and 1/1 uncertain. The corresponding totals for Hoh were 223/54, 6/2, 28/10, 78/26 and 5/3. In addition to sporocarps, we analyzed 10 samples each of leaves, twigs and bole wood from EcM trees (one sample of wood was consumed during a malfunction of the mass spectrometer), 10 samples of CWD, 15 samples each of O-horizon and 0–10 cm mineral soil, and three samples of 10–20 cm mineral soil at both DP and Hoh. Mean δ15N and δ13C for each pool, by area, are given in Table 1.

Table 1.  δ15N and δ13C of macrofungus, plant and soil samples collected at lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA
Comparisonn (DP, Hoh)δ15Nδ13C
DPHohPDPHohP
  • Values are means ± SD in ‰.

  • EcM, ectomycorrhizal; Sap, saprotrophic; SapH, saprotrophic on soil (‘humus’); SapL, saprotrophic on litter; SapW, saprotrophic on wood; CWD, coarse woody debris; Ohor, soil O-horizon; Mhor, upper mineral soil.

  • Most comparisons made with t-tests for independent groups; species of fungi in common compared with paired t-test. For fungi, n = number of species. For plant and soil pools, n = number of samples. P values significant at α= 0.05 in bold type; ***, P < 0.001.

  • Species with ≥ 3 analyses at both DP and Hoh: Cantharellus formosus, Chroogomphus tomentosus, Cortinarius gentilis, Cortinarius montanus, Gymnopus acervatus, Laccaria bicolor, Pholiota decorata, Rhodocollybia extuberans, Russula brevipes, Russula fragilis, Tricholoma saponaceum, Xeromphalina cornui.

  • During analysis of wood samples, one sample of Pseudotsuga menziesii was lost during an equipment malfunction.

Fungi
 All species89, 95 3.7 ± 4.1 1.8 ± 4.50.003−24.9 ± 1.3−24.3 ± 1.50.004
 EcM65, 54 5.5 ± 3.2 4.7 ± 3.80.179−25.4 ± 0.8−25.2 ± 0.70.160
 Sap23, 38−1.2 ± 1.7−2.3 ± 1.50.006−23.3 ± 1.4−22.9 ± 1.20.235
 SapH 3, 2 1.0 ± 1.5−0.4 ± 1.30.378−22.1 ± 1.3−23.5 ± 0.10.215
 SapL 7, 10−2.8 ± 1.4−2.8 ± 1.00.995−24.5 ± 1.1−23.6 ± 1.10.098
 SapW13, 26−0.8 ± 0.9−2.3 ± 1.50.003–23.0 ± 1.2−22.6 ± 1.20.383
 Cortinarius17, 7 5.8 ± 2.4 6.1 ± 1.60.784−25.5 ± 0.9−25.3 ± 0.60.566
 Lactarius 5, 5 4.3 ± 1.1 3.0 ± 2.10.257−25.1 ± 0.5−25.1 ± 0.50.965
 Russula 6, 8 2.7 ± 1.5 1.7 ± 2.90.475−25.6 ± 0.8−25.6 ± 1.10.964
 Species in common12 1.8 ± 3.9 1.6 ± 4.30.747−24.8 ± 1.6–24.6 ± 1.30.494
EcM trees
 Leaves10, 10−5.8 ± 1.8−3.7 ± 0.50.003–30.2 ± 0.7−30.5 ± 1.40.629
 Twigs10, 10−5.6 ± 1.1−4.1 ± 0.70.002–28.8 ± 0.8−29.0 ± 1.60.695
 Bole wood 9, 10 0.9 ± 1.6 2.6 ± 1.90.051−25.9 ± 1.2−25.9 ± 1.00.949
CWD10, 10 0.0 ± 0.7−0.6 ± 1.40.233−26.5 ± 1.0−25.4 ± 0.80.009
Ohor15, 15 0.8 ± 0.7−1.5 ± 0.5***−28.4 ± 0.5−27.4 ± 0.3***
Mhor (0–10 cm)15, 15 4.3 ± 0.4 2.1 ± 0.8***−26.8 ± 0.2−26.0 ± 0.2***
Mhor (10–20 cm) 3, 3 4.7 ± 0.5 2.9 ± 0.40.004−26.6 ± 0.3−25.7 ± 0.20.004

More support for the EcM–Sap divide

Sporocarps of EcM and saprotrophic fungi differ in δ15N and δ13C (Gebauer & Taylor, 1999; Hobbie et al., 1999a, 2001; Kohzu et al., 1999; Henn & Chapela, 2001; Taylor et al., 2003), and dual isotope plots have shown a clear separation between the two groups (Kohzu et al., 1999; Henn & Chapela, 2001; Hobbie et al., 2001; Taylor et al., 2003). The line separating the two groups in plots of δ13C vs δ15N has been termed the ‘EcM–Sap divide’ (Henn & Chapela, 2001). Although some of the previously reported data sets contained considerable overlap of EcM and saprotrophic taxa (data points appearing on the ‘wrong’ side of the divide), at least some of this imprecision can be attributed to small sample size, incomplete or questionable identifications, and pooling of data from different sites.

At DP and Hoh, the mean δ15N and δ13C values of EcM and saprotrophic fungi differed significantly, whether the areas were considered separately or together (P < 0.001 for both isotopes at both areas in both cases). The distinction between the groups is clear when δ13C is plotted against δ15N, whether as individual samples (not shown) or species means (Fig. 1). Previously, attempts have been made to assign species to the EcM or saprotroph groups based on a single isotope. For instance, Hobbie et al. (2001) suggested that, for their data set, δ13C was a better indicator of trophic type than δ15N, and that δ13C = −24‰ was an effective delimiter of EcM (< −24‰) and saprotrophic (> −24‰) taxa. Taylor et al. (2003) also emphasized δ13C in considering Chalciporus piperatus to be a saprotroph, despite it having a rather high δ15N (δ13C = −22.6 to −21.4‰; δ15N = 8.3–9.4‰) compared with the mean for EcM species in their study (6.2‰). However, for our study areas there is no single value of δ13C or δ15N that separates all EcM fungi from all saprotrophic fungi (Fig. 1). Thus we agree with Hobbie et al. (2001) that a combined index provides the best means of distinguishing the two groups.

Figure 1.

Nitrogen and carbon stable isotope values for sporocarps of ectomycorrhizal and saprotrophic macrofungi collected at (a) lower Deer Park Road (DP) and (b) Hoh River Valley (Hoh), Olympic National Park, Washington, USA. Data points represent species means. Filled circles, ectomycorrhizal fungi; open triangles, saprotrophic fungi on soil (‘humus’); open squares, saprotrophic fungi on plant litter; open circles, saprotrophic fungi on wood. Species means and n are provided in Appendix 1.

Within the saprotrophic fungi, there were differences in mean δ15N and δ13C values among SapH, SapL and SapW at both DP and Hoh (Table 2). However, in most cases the differences were not statistically significant, and overlap among the three groups was considerable (Fig. 1). Contrary to findings in previous studies that SapL fungi were enriched in 15N compared with SapW fungi (Gebauer & Taylor, 1999; Kohzu et al., 1999), we found SapL fungi to be depleted in 15N compared with SapW fungi, both at DP and Hoh. The difference was statistically significant at DP, but not at Hoh (Table 2). Thus it appears that the pattern of δ15N among different groups of saprotrophic macrofungi can vary from site to site.

Table 2.  Multiple-comparison test summary for δ15N and δ13C in saprotrophic macrofungi from lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA
Trophic type (n)δ15N (‰)δ13C (‰)SapHSapLSapW
  1. δ15N and δ13C values are means. SapH, saprotrophic on soil (‘humus’); SapL, saprotrophic on litter; SapW, saprotrophic on wood. Comparisons made with anova–Tukey hsd for unequal n. P values for δ15N listed on lower left, below the diagonal; those for δ13C listed on upper right, above the diagonal. n = number of species. P values significant at α= 0.05 in bold type.

DP
SapH (3) 1.0−22.10.0410.595
SapL (7)−2.8−24.50.0020.052
SapW (13)−0.8−23.00.1710.011
Hoh
SapH (2)−0.4−23.50.9990.720
SapL (10)−2.8−23.60.2200.172
SapW (26)−2.3−22.60.3820.708

In nearly all cases the isotope data were consistent with the trophic-type designations based on the literature. In cases where the literature information was equivocal, the isotope data strongly supported EcM status (Phaeocollybia spp.); moderately supported EcM status (Clavulina cristata, Entoloma nitidum); or provided little clarification (Bondarzewia mesenterica, Phylloporus rhodoxanthus). Although belonging to a genus thought to be entirely EcM, Chroogomphus tomentosus had an uncharacteristically low δ15N at DP (−4.4‰, n = 10). In fact its δ15N was the lowest of any species at DP, including the saprotrophic taxa. Its value at Hoh was −0.7‰ (n = 3), near the low end of the typical range for EcM fungi. Kohzu et al. (1999) reported three δ15N analyses of C. tomentosus from subalpine conifer forests in Japan. One analysis was similar to the mean value observed at DP (−3.5‰), while the others were typical of EcM fungi (4.1 and 4.6‰). Other species of Chroogomphus have been reported to associate with mycorrhizas of Suillus and Rhizopogon species (Agerer, 1990), suggesting that they might be mycoparasitic. If true (and direct evidence is lacking), that might explain the unusual δ15N values. However, the δ15N of mycoheterotrophic plants from DP and Hoh that are parasitic on EcM fungi is higher than that of their hosts (Trudell et al., 2003), not lower, as would be the case here (δ15N for Suillus lakei and Suillus punctatipes are 5.8 and 11.6‰, respectively). This suggests that if C. tomentosus is mycoparasitic, the mechanism(s) by which it obtains N from its host must be different from that of mycoheterotrophic plants and, perhaps, be similar to that of typical EcM phytobionts. Only one species (Cortinarius variosimilis) yielded a dual isotope signature that clearly placed it in the ‘wrong’ trophic group. It is generally believed that all species in Cortinarius are EcM. However, the values for C. variosimilis15N = −1.5‰, δ13C = −23.1‰, n = 2) placed it well within the saprotroph group (Fig. 1a). Kohzu et al. (1999) reported somewhat similar values (δ15N = 0.2‰, δ13C = −22.6‰) for an unidentified species of Cortinarius from a mixed forest in Japan. Otherwise, δ15N and δ13C values for Cortinarius reported here and in the literature have been much higher and much lower, respectively, consistent with the genus being EcM. The reasons for the occasional contrary results are not clear, but they argue for caution in attributing particular ecophysiological functions to taxa above the species level.

Taxonomic patterns

Some early reports of δ15N values in EcM fungus taxa (e.g. Lilleskov et al., 1997; Hobbie et al., 1999a) suggested there were significant differences among genera. However, sample sizes were relatively small and many of the samples were identified only to genus, making it difficult to determine the significance of the results. None of the early studies included sufficient intraspecies replication to allow possible differences among species in a genus to be assessed. In a preliminary report from this study, we showed significant differences among genera and congeneric species of EcM fungi (Trudell et al., 2001). More recently, Taylor et al. (2003) reported similar findings based on a large data set from two sites in Sweden. Although Taylor et al. (2003) also reported significant differences at the family level, we agree with their decision not to emphasize them. Families such as Tricholomataceae and Cortinariaceae contain large numbers of saprotrophic, as well as EcM, taxa and it is not clear that the families of macrofungi, as currently circumscribed, represent entirely natural groupings.

At both DP and Hoh, we found statistically significant differences in δ15N and δ13C among both genera (Table 3) and species. Based on δ15N, most genera fall into high (> 6‰) and low (< 3‰) groups (Fig. 2), with Lactarius at DP slightly exceeding 4‰. A similar pattern appears in the data of Taylor et al. (2003; their Figure 2). Mean δ15N of all genera are either < c. 4‰ or > 6‰, with the exception of Lactarius (c. 5‰). Occurrence as a ‘high’ or ‘low’ genus is fully consistent between the two studies. Although the genera also differ in δ13C, the values are spread more or less evenly between −26.0 and −24.5‰ in both studies.

Table 3.  Multiple-comparison test summary for δ15N in 10 ectomycorrhizal genera from lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA
Genusδ15NDP (n)δ15NHoh (n)AmanBoleCortHydnHygrInocLactPhaeRussTric
  1. δ15N values are means of species means, in ‰. Comparisons made with anova–Tukey hsd for unequal n. P values and codes for DP are listed on lower left, below the diagonal; those for Hoh are listed on upper right, above the diagonal. Only genera with ≥ 3 species having ≥ 3 analyses at an area are included. Not all cells are filled because some genera were not present at one or the other area in sufficient abundance to be included in the analysis. Full genus names are given in far-left column. n = number of species. P values significant at α= 0.05 in bold type; ***, P < 0.001.

Amanita1.5 (4)0.0760.0831.0000.9970.0320.998
Boletus6.9 (3)0.9990.0980.1991.0000.184
Cortinarius6.7 (10)6.1 (6)0.2220.2460.9760.081
Hydnellum7.4 (3)0.995
Hygrophorus2.5 (3)0.0770.024
Inocybe1.7 (3)1.0000.0431.000
Lactarius4.3 (5)2.4 (4)0.2970.2790.8270.0941.000
Phaeocollybia7.7 (3)0.086
Russula2.8 (5)2.3 (6)0.0190.0351.0000.752
Tricholoma8.5 (6)0.5060.9770.0040.010***
Figure 2.

Variation in nitrogen and carbon stable isotope values in genera of ectomycorrhizal macrofungi from (a) lower Deer Park Road (DP, filled circles) and (b) Hoh River Valley (Hoh, open circles), Olympic National Park, Washington, USA. Data points represent means of species means; error bars, SE. Numbers in brackets indicate number of species analyzed.

Within genera having ≥ 3 species with ≥ 3 analyses, individual species also exhibited distinct signatures (Figs 3–5), except in Amanita (at Hoh; not shown). In most cases, groupings of species or correlations with infrageneric taxonomy were not obvious. However, within the EcM fungi, species in some genera exhibited greater variation in δ15N than those in most other genera. For instance, Hygrophorus species (Fig. 4c) showed much larger variation than did Boletus species (Fig. 4a), despite having similar sample sizes. Several species of Tricholoma also showed large variation (Fig. 4f) compared with species in other genera (Figs 3, 4b,d,e). Conceivably, species with highly variable signatures could be accessing a relatively wide range of N sources, or could be more physiologically diverse compared with species with less variable signatures. However, the species of Hygrophorus and Tricholoma in the Taylor et al. (2003) study did not show particularly large variation, so the significance of our observations is not clear. Although the isotope signatures of saprotrophic fungi have not received as much attention as those of EcM fungi, our data for Hypholoma and Pholiota (Fig. 5) suggest that the patterns of saprotrophic species can be as distinct as those of EcM species. Thus our findings are consistent with those of Taylor et al. (2003), and support their conclusion that the species is the most informative taxonomic level. This reinforces the need for species-level identification and replication of samples in future studies involving sporocarp isotope abundances.

Figure 3.

Variation in nitrogen and carbon stable isotope values in species of ectomycorrhizal macrofungi in three genera from lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA. Data points represent means of analyses; error bars, SE. Numbers in brackets indicate number of analyses. (a) Cortinarius at DP; (b) Cortinarius at Hoh; (c) Lactarius at DP; (d) Lactarius at Hoh; (e) Russula at DP; (f) Russula at Hoh. These are the three genera with sufficient samples to conduct between-area comparisons (see Table 1 for sample size data).

Figure 4.

Variation in nitrogen and carbon stable isotope values in species of ectomycorrhizal macrofungi in six genera from lower Deer Park Road (DP, filled circles) and Hoh River Valley (Hoh, open circles), Olympic National Park, Washington, USA. Data points represent means of analyses; error bars, SE. Numbers in brackets indicate number of analyses. (a) Boletus; (b) Hydnellum; (c) Hygrophorus; (d) Inocybe; (e) Phaeocollybia; (f) Tricholoma.

Figure 5.

Variation in nitrogen and carbon stable isotope values in species of saprotrophic macrofungi in two genera from lower Deer Park Road (DP, filled circles) and Hoh River Valley (Hoh, open circles), Olympic National Park, Washington, USA. Data points represent means of analyses; error bars, SE. Numbers in brackets indicate number of analyses. (a) Hypholoma; (b) Pholiota.

Effect of macrofungus species composition on spatial patterns

Previously, we showed there were striking differences in the macrofungi at DP and Hoh (Trudell & Edmonds, 2004). At DP, EcM species in genera such as Cortinarius, Hydnellum, Sarcodon, Suillus and Tricholoma were dominant. At Hoh, the dominant EcM species belonged to different genera, including Amanita, Boletus, Inocybe, Phaeocollybia and Russula, and saprotrophic fungi accounted for a significantly greater proportion of the macrofungi. These differences are an important factor behind the significantly higher overall δ15N of macrofungi at DP compared with Hoh, whether considering species means (Table 1) or all samples (4.5‰ at DP vs 1.9‰ at Hoh, P < 0.001). First, the significantly higher proportion of EcM taxa, which tend to be enriched in 15N, at DP (69 vs 62% based on species; 86 vs 72% based on collections) leads to a higher overall value there. The greater dominance by EcM fungi at DP also leads to a slightly lower δ13C there. Second, among the EcM fungi, a large proportion of the species at DP belong to high-δ15N genera including Cortinarius, Hydnellum, Suillus and Tricholoma (Trudell & Edmonds, 2004; their Figure 1). Although high-δ15N taxa also occur at Hoh (e.g. species of Boletus, Cortinarius, Phaeocollybia and Tricholoma), they comprise a much smaller portion of the community than do species in low-δ15N genera such as Amanita, Inocybe and Russula.

When compared by trophic group, statistically significant differences exist only for δ15N in saprotrophs (SapH-L-W combined) and SapW. When compared by genus (those having ≥ 3 species with ≥ 3 analyses at both areas: Cortinarius, Lactarius and Russula), there are no statistically significant differences. Comparison based on the 12 species with ≥ 3 analyses at both areas likewise showed no statistically significant difference in δ15N or δ13C. This pattern of results suggests that the between-area differences were caused more by a difference in the species present than by direct environmental effects. A similar conclusion was drawn by Taylor et al. (2003) when considering differences between their two study areas and among plots at one of the areas.

A species effect also is clear in the data reported by Gebauer and Taylor (1999). They observed considerable overlap in δ15N between EcM and saprotrophic (mostly SapH) species. However, the range in their data was very narrow (6‰; −3 to 3‰), principally because there were no species from high-δ15N genera. Instead, their EcM taxa were from characteristically low- to moderate-δ15N genera such as Amanita, Laccaria, Russula and Xerocomus. Conversely, Lilleskov et al. (2002) reported that four species and two genera of EcM fungi exhibited higher δ15N values at high-N sites than at nearby lower-N sites, suggesting that, in some cases, environmental effects might be greater than species effects.

We also compared the DP and Hoh areas with Åheden and Stadsskogen, Sweden (Taylor et al., 2003), focusing on the EcM taxa because of the relatively small number of saprotrophs at the Swedish sites. When the four areas were compared using all species encountered at each, DP and Hoh differed significantly from Åheden in δ15N (Table 4). With respect to δ13C, Hoh and Stadsskogen differed, although the result was not quite significant statistically (P = 0.059). However, when DP and Hoh were compared with Åheden and Stadsskogen on the basis of shared species, there were no statistically significant differences between US–Sweden pairs in the δ15N or δ13C of the fungi, and correlations in species δ15N values between sites were high (Table 5). This also suggests a strong effect of species composition. Such comparisons necessarily are limited to sites with similar environmental conditions and assemblages of macrofungi. As environmental conditions, such as vegetation, temperature and soil moisture, become increasingly different, N- and C-cycling processes will differ and there will be fewer macrofungi in common between sites. For instance, Griffith et al. (2002) presented 30 δ15N/δ13C analyses of fungi collected from a grassland at Sourhope, UK. Species of Hygrocybe and clavarioid fungi, all of which are almost certain to be saprotrophs, yielded δ15N (c. 8–18‰) and δ13C values (< −27‰) that, if plotted with our data (Fig. 1), would place them among the EcM taxa, well separated from the saprotrophs. No data for plants or soils were presented by Griffith et al. (2002), and only one of the fungi (Cystoderma amianthinum) has been included in previous studies. Thus it is difficult to determine the cause(s) for the results, although differences in N dynamics between grasslands and forests seem likely to be an important factor. Nevertheless, this reinforces the need to include as many pools as possible when making between-site comparisons and to exercise caution when interpreting results and inferring causes.

Table 4.  Multiple-comparison test summary for between-area comparisons of δ15N and δ13C in sporocarps of ectomycorrhizal fungi from four areas – lower Deer Park Road (DP) and Hoh River Valley (Hoh); Olympic National Park, Washington, USA; and Åheden and Stadsskogen, northern and central Sweden, respectively
Site (n)δ15N (‰)δ13C (‰)DPHohÅhedenStadsskogen
  1. Data for Swedish sites from Taylor et al. (2003). Values for δ15N and δ13C are means. Comparisons made with anova–Tukey hsd for unequal n.

  2. P values for δ15N are listed on lower left below the diagonal, those for δ13C are listed on upper right, above the diagonal. n = number of species. P values significant at α= 0.05 in bold type.

DP (65)5.5−25.40.7710.5740.329
Hoh (54)4.7−25.20.5490.1930.059
Åheden (28)7.8−25.80.0500.0020.999
Stadsskogen (109)5.8−25.80.9570.2800.116
Table 5.  Summary of between-area comparisons of δ15N and δ13C in sporocarps of ectomycorrhizal fungi from four areas – lower Deer Park Road (DP) and Hoh River Valley (Hoh); Olympic National Park, Washington, USA; and Åheden and Stadsskogen, northern and central Sweden, respectively
Comparison (n)δ15Nδ13C
tPrPtPrP
  • Data for Swedish sites from Taylor et al. (2003). Comparisons made with paired t-tests and Pearson's product-moment correlation (r). n = number of species. P values significant at α= 0.05 in bold type.

  • Species included in the analyses are: Amanita fulva (nDP = 0, nHoh = 5, nÅh = 0, nStad = 6), Boletus edulis (0-1-0-5), Cortinarius brunneus (5-0-0-2), Cortinarius gentilis (5-3-0-4), Cortinarius laniger (19-0-6-2), Cortinarius malachius (0-3-2-3), Cortinarius muscigenus (10-0-0-3), Cortinarius semisanguineus (4-0-3-7), Cortinarius vibratilis (5-0-0-7), Craterellus (or Cantharellus) tubaeformis (5-0-0-4), Entoloma nitidum (considered a saprotroph by Taylor et al. (2003); 0-3-0-3), Gymnopilus penetrans (0-3-1-0), Hebeloma mesophaeum (4-0-0-2), Hydnellum peckii (10-0-0-1), Hygrophoropsis aurantiaca (0-5-0-2), Hygrophorus camarophyllus (2-0-0-5), Hygrophorus olivaceoalbus (0-5-0-5), Laccaria bicolor (4-5-2-0), Paxillus atrotomentosus (2-0-0-1), Russula queletii (0-2-0-1), Russula xerampelina (5-0-0-1), Tricholoma flavovirens (2-0-4-2), Tricholoma vaccinum (2-0-0-1), Tricholoma virgatum (4-1-1-2).

DP-Åheden (5)−0.6970.5240.860.060−0.3250.762−0.470.430
DP-Stadsskogen (15) 0.1390.8910.690.005−0.0810.937 0.310.258
Hoh-Åheden (4)−2.0410.1340.960.045−0.2450.822 0.610.392
Hoh-Stadsskogen (9)−1.0440.3270.710.034 0.7330.484 0.720.028

Ecophysiological basis for species differences

The ecophysiological basis for the differences in δ15N among EcM species (and saprotrophic species too, although they have received much less attention) is not well understood. However, as data from field and laboratory studies accumulate, interesting patterns are emerging that suggest possible explanations. It is important to keep in mind that natural abundance stable isotope ratios present a time-integrated picture of the isotope effects of multiple processes (Robinson, 2001). Thus a given δ15N value could be produced by more than one set of processes. The pattern of overlapping species values arrayed along a continuum (Figs 3–5) suggests interaction of multiple factors.

The ecophysiological factor for which there is strongest support is the ability to utilize organic forms of N. Taylor et al. (2000) described variation among EcM taxa in organic N utilization, and showed how the relative abundance of organic N users correlated with soil mineral N abundance along a transect from central to northern Europe (Erland & Taylor, 2002). Lilleskov et al. (2002) observed a similar pattern along a local N-deposition gradient associated with a fertilizer manufacturing facility in Alaska, USA. Their combination of culture studies with isotope analyses of field-collected sporocarps showed a positive relationship between δ15N and organic N use, and a strong negative correlation between the occurrence of organic N users and soil mineral N concentration. In a recent culture study, Henn and Chapela (2004) reported a correlation between mycelium δ15N and the amount of ammonium-N incorporated into biomass, and suggested that their observation could provide a mechanism by which fungi that utilize organic N become enriched in 15N relative to those that utilize inorganic N. Although we have no site-specific data on organic N use by the taxa at DP and Hoh, species composition at the two areas relative to N availability is consistent with organic N use being a key factor. Putative organic N users, such as species in Cortinarius, Suillus and Tricholoma, are much more abundant at DP than at Hoh, where the N content of fungi, plants and soils and the soil mineral N concentrations are higher (Trudell & Edmonds, 2004). These taxa exhibited moderate to very high δ15N values.

The EcM macrofungi at DP and Hoh also appear to differ in mycelial morphology (Trudell & Edmonds, 2004). The dominant genera at DP (Cortinarius, Hydnellum, Suillus, Tricholoma) contain high proportions of species that form extensive extraradical mycelia, whereas the dominant genera at Hoh (Amanita, Inocybe, Lactarius, Russula) contain high proportions of species that form diffuse, spatially limited extraradical mycelia. These characteristics also appear to be associated with different reproductive and colonization strategies (Taylor et al., 2000, 2003; Erland & Taylor, 2002). The extensive mycelia are thought to become established infrequently, often by vegetative means, and to be longer-lived, whereas the diffuse mycelia become established more frequently, usually by spores, but do not persist. Fungi are flexible with respect to N metabolism, and are thought to store N in a variety of insoluble forms such as protein inclusions and Woronin bodies (Jennings, 1995). The mycelia of saprotrophic basidiomycetes have been shown to remobilize and translocate biomass and stored mineral nutrients (Boddy, 1999), and it is likely that many EcM fungi function similarly. As the degree of metabolic processing of N increases, so does the δ15N of residual N (Peterson & Fry, 1987; Yoneyama, 1996). Because large, long-lived mycelia offer greater potential for repeated metabolic processing of assimilated N than do small, short-lived ones, the former would be expected to exhibit higher δ15N. This is the pattern that appears at our sites. Species of Cortinarius, Hydnellum, Suillus and Tricholoma exhibit higher δ15N values than Amanita, Inocybe, Laccaria and most species of Russula, consistent with their comparative mycelial morphology (Agerer, 2001). Lilleskov et al. (2002) also reported high δ15N values in presumably long-lived rhizomorph-forming species relative to nonrhizomorph-formers. The observation by Henn and Chapela (2004) that EcM and saprotrophic fungi grown in liquid culture appeared to return isotopically depleted (c.−3‰) ammonium to the medium suggests one possible mechanism for enrichment of long-lived mycelia. The authors proposed that life history and territorial behavior of mycelia might play an important role in determining the δ15N of the mycelium and associated ecosystem components.

It also has been suggested that EcM fungi that transfer a relatively low proportion of their assimilated N to associated phytobionts should exhibit higher δ15N values (Hobbie et al., 2000; Hobbie & Colpaert, 2003). Recent support for this proposition comes from Hobbie and Colpaert (2003) who observed that, in culture, Suillus luteus transferred less N to Scots pine seedlings than did Thelephora terrestris, and exhibited higher δ15N. Interaction of these factors – organic N use, degree of metabolic processing, and proportion of N transferred to phytobionts – plus others that are likely to exist would be expected to produce a range of δ15N values depending on the relative importance of the individual factors. For instance, a species that is restricted to inorganic N, does relatively little metabolic processing, and passes a high proportion of its N to phytobionts or the soil could have a very low δ15N, whereas a species that uses organic N, does extensive metabolic processing, and passes a low proportion of its N to phytobionts or the soil could have a very high δ15N. Many intermediate values could result from combinations of these factors. However, the possibility that there are high- and low-δ15N genera (Fig. 2) suggests that these features might occur as syndromes, reducing the number of intermediate states.

Taylor et al. (2000) discussed organic N use by EcM fungi in the context of R (ruderal), C (competitive) and S (stress-tolerant) life-history patterns (Grime, 1977). They pointed out that the ability to access organic N in a low-N-availability habitat is consistent with the S strategy, whereas reliance on mineral N and inability to utilize organic N is consistent with the R strategy. The observations on mycelial morphology and N transfer also fit well within this framework. Development of a large, long-lived mycelium and high retention of N in low-nutrient habitats would represent stress-tolerance characteristics. Development of small, ephemeral mycelia and low N retention would represent ruderal characteristics. In all three cases, species with presumed S-type characteristics exhibit high δ15N values, and species with presumed R-type characteristics exhibit low δ15N values. Thus the R-C-S life-history patterns appear to offer a useful conceptual framework to guide continuing research into the ecophysiological basis of stable isotope patterns in macrofungi.

The generally observed pattern of δ15N increasing with depth in soils (Nadelhoffer & Fry, 1988) suggests that this also could be a factor in the δ15N of soil fungi. Our lack of knowledge of the depth distribution of the mycelium of most fungi prevents this hypothesis from being tested rigorously. However, at Hoh several species of Phaeocollybia occur, the stipes of which extend downward as long pseudorhizas, often into the mineral soil. The 0–10 cm mineral soil at Hoh was considerably enriched relative to the O-horizon, and the 10–20 cm mineral soil was enriched even further (Table 1). The δ15N values for Phaeocollybia spp. (Fig. 4e; Appendix 1) are high, suggesting that they could be accessing enriched N from the mineral soil. Occurrence of a ‘rooting-depth’ effect is also suggested by our data for three herbaceous plant species at Hoh. Trillium ovatum, whose roots occurred in mineral soil, exhibited a higher δ15N (−1.11 ± 0.72; n = 5) than Maianthemum dilatatum (−3.91 ± 0.73; n = 5) and Oxalis oregana (−4.35 ± 0.30; n = 5), both of whose rhizomes and roots were in the organic horizon. Molecular methods for determining the vertical distribution of fungi within soil profiles are being developed (Dickie et al., 2002; Kuyper & Landeweert, 2002; Landeweert et al., 2003; Rosling et al., 2003), and their availability should allow the relationships between mycelium depth and δ15N to be assessed more fully.

The basis for δ13C values in macrofungus sporocarps has received less attention than that for δ15N. It has been suggested that δ13C reflects the C source more faithfully than δ15N does the N source (Hobbie et al., 2001), due to fewer isotope effects during metabolic processing, but data are too few to determine whether this is so. While studies with fungi have shown that substantial fractionation can occur during C metabolism (Abraham et al., 1998; Henn & Chapela, 2000, 2001; Henn et al., 2002), the magnitude of fractionation appears to be similar across taxa. For instance, Högberg et al. (1999b) showed that differences in δ13C values among EcM sporocarps reflected specific phytobiont sources (pine vs spruce vs birch and other angiosperms), and that generalist fungi received most of their C from the upper-canopy trees (predominantly pines). Unfortunately, patterns of specificity in mycobiont–phytobiont associations for North American fungi are not well documented. Therefore it is currently not possible to assess the degree to which the δ13C values of the EcM fungi at DP and Hoh reflect phytobiont associations.

Ecosystem patterns

Taken together, the isotope signatures of the fungi, plants and soils at DP and Hoh exhibited many of the patterns seen in other data sets (Fig. 6). For instance: (1) δ15N in EcM fungi was comparable to that in mineral soil and greater than that in plant foliage and saprotrophic fungi; (2) δ13C in both EcM and saprotrophic fungi was greater than that in their bulk substrates as was δ15N in EcM fungi; (3) δ15N and δ13C in soils both increased with depth; (4) δ13C in EcM fungi was less than that in saprotrophic fungi, but greater than that in plant foliage. In addition to these overall patterns, the observed magnitudes of between-pool differences in δ15N and δ13C generally fell within the range of reported values (Table 6).

Figure 6.

Nitrogen and carbon stable isotope values for 10 ecosystem pools at (a) lower Deer Park Road (DP) and (b) Hoh River Valley (Hoh), Olympic National Park, Washington, USA. Data points represent means; error bars, SE. n for each pool is given in Table 1. CWD = coarse woody debris; EcMF = ectomycorrhizal fungi; Mhor = upper mineral soil; Ohor = soil O-horizon; SapH = saprotrophic fungi on soil (‘humus’); SapL = saprotrophic fungi on plant litter; SapW = saprotrophic fungi on wood.

Table 6.  Differences in δ15N and δ13C between pairs of ecosystem pools at lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA
Poolsδ15N (‰)δ13C (‰)
DPHohLiteratureDPHohLiterature
EcMF-SapF  6.7 7.01–10−2.1−2.3−3 → –2
EcMF-Leaves 11.3 8.44–12 4.8 5.3 1–5
EcMF-Wood  4.6 2.05.1 0.5 0.8 1.4
EcMF-Ohor  4.7 6.22–7 3.0 2.2 1–4
EcMF-Mhor  1.2 2.6−4 → –2 1.4 0.8∼0
SapH-Ohor  0.2 1.12.5 6.4 3.9
SapH-Mhor −3.4−2.5 4.7 2.5
SapL-Leaves  3.0 0.9 5.7 6.9 4
SapL-Twigs  2.8 1.3 4.3 5.5
SapL-Ohor −3.6−1.3−3–0 3.9 3.8 3–4
SapW-Wood −1.7−5.0−3–0 2.9 3.3 3–4
SapW-CWD −0.8−1.7 3.5 2.7 4.3
Leaves-Twigs −0.2 0.4−1.4−1.4
Leaves-Wood −6.7−6.4−4.3−4.6
Leaves-Ohor −6.0−2.2−4 → –1−1.8−3.1−2 → –1
Leaves-Mhor−10.2−5.8−10 → –3−3.5−4.5−3 → –2
Twigs-Wood −6.5−6.7−2.9−3.2
Wood-CWD  0.9 3.2 0.6−0.5
Ohor-Mhor −3.6−3.6−6 → –2−1.6−1.4−2 → –1

Mean δ15N values in fungi (by trophic type) and soils at DP all were greater than those of the corresponding pools at Hoh, with the exception of SapL fungi, in which the two means were essentially the same (Table 1). Values for nonEcM plant pools also were higher at DP (data not shown). Conversely, mean δ15N values in EcM plant pools all were higher at Hoh, thus producing smaller ΔEcM fungi–foliage values there, compared with DP. It has been suggested that δ15N differences between EcM fungi and EcM plant foliage (Hobbie et al., 2000); between EcM plant roots and soil (Högberg et al., 1996); and between foliage and soil (Garten & van Miegroet, 1994; Emmett et al., 1998; Hobbie et al., 2000) could be effective indicators of a site's N status, with larger differences being produced when mineral N is less available. Hobbie and Colpaert (2003) presented a model that suggests a mechanistic basis for the EcM fungi–foliage relationship reported by Hobbie et al. (2000), conducted a culture study, and used the results to assess the model. The authors suggested that the low δ15N values of EcM plant foliage and high values in EcM fungi in low-productivity, N-limited environments result in part from high retention of 15N-enriched N by EcM fungi, this retention being driven by an increased flux of C to the fungi in response to the nutrient limitation. If this is so, then the δ15N patterns of plants might provide an indication of C allocation to EcM fungi (Hobbie & Colpaert, 2003). However, contrary results were reported by Lilleskov et al. (2002) who found that, along a gradient of increasing mineral N availability, δ15N values of EcM fungi increased, while those of spruce foliage decreased.

Our data are consistent with Hobbie and Colpaert's (2003) model and interpretation. ΔEcM fungi–foliage is greater at DP than Hoh and, among all pools, only the EcM plants have lower δ15N at DP than at Hoh. Earlier (Trudell & Edmonds, 2004), we showed that N availability is greater at Hoh and that EcM fungus taxa with presumably high C demands (based on mycelial morphology and abundance of sporocarps) are more abundant at DP, as would be predicted by the model. The reasons for the apparent disagreement between the model and the findings of Lilleskov et al. (2002) remain unclear.

Conclusions

Our data set, derived from two old-growth, temperate conifer forests, extends previous observations and provides additional support for the existence of distinctive patterns of δ15N and δ13C in northern conifer forests. The mean δ15N and δ13C values of EcM and saprotrophic fungi differed greatly in both forests, and a dual 15N/13C isotope approach provided the best means of distinguishing the two groups. Within both EcM and saprotrophic groups, δ15N and δ13C differed among genera and species, and the difference in species composition was an important determinant of the different overall δ15N of the EcM fungi at DP and Hoh. Patterns among fungi and other ecosystem pools were similar to those observed elsewhere, with EcM fungi and mineral soil exhibiting relatively high δ15N and moderate δ13C values, saprotrophic fungi exhibiting low to moderate δ15N and high δ13C values, and EcM plant foliage exhibiting the lowest δ15N and δ13C values.

Simultaneous variation in multiple ecophysiological traits such as organic N use, mycelial morphology, and proportional transfer of N to phytobionts appears to underlie the pattern of variation in the isotope signatures of EcM fungi. The wide range of dual isotope patterns suggests a high degree of functional diversity among these fungi. Ecosystem δ15N patterns at DP and Hoh are consistent with the model of Hobbie and Colpaert (2003), which suggests that these patterns are influenced strongly by soil N availability and C allocation within the EcM symbioses. Thus fungus-plant δ15N differences might prove to be an effective indicator of site N status.

Acknowledgements

We thank Olympic National Park for allowing access to the study sites; Joe Ammirati and Michelle Seidl (Cortinarius), Brandon Matheny (Inocybe), Lorelei Norvell (Phaeocollybia), Clarke Ovrebo (Tricholoma), and Ben Woo (Russula) for identification assistance; Joe Ammirati for updating species authorities; Bill Griffis, Paul Brooks, Dongsen Xue and Julia Cox for laboratory analyses; and Joe Ammirati, Charlie Halpern, Tom Hinckley, Erik Lilleskov, and three anonymous reviewers for thoughtful comments on earlier versions of the manuscript. The study was supported by an EPA NNEMS fellowship and University of Washington College of Forest Resources Lockwood fellowship to S.A.T., and grants from the Global Forest Society, Mycological Society of America, North American Mycological Association, and Northwest Scientific Association. This is Global Forest contribution GF-18-1999-31. The manuscript submitted was subjected to EPA's peer and administrative reviews and was approved for publication. Mention of trade names and commercial products does not constitute endorsement or recommendation of use.

Appendix

Table Appendix1.  δ15N and δ13C of sporocarps of ectomycorrhizal and saprotrophic macrofungi collected at lower Deer Park Road (DP) and Hoh River Valley (Hoh), Olympic National Park, Washington, USA
SpeciesTypeAreanδ15Nδ13C
  • Values are means ± SD, in ‰. n = number of samples analysed.

  • Trophic types: EcM, ectomycorrhizal; SapH, saprotrophic on soil; SapL, saprotrophic on plant litter; SapW, saprotrophic on wood; Par, parasite. Parasites were included with SapW in statistical calculations.

  • Interpreted in broad sense.

  • §

    Xerocomus chrysenteron (Bull.) Quél.

  • Xerocomus truncatus Singer, Snell & E.A. Dick in Snell.

  • **

    Xerocomus zelleri (Murrill) Snell.

  • ††

    Clavulina coralloides (L.: Fr.) J. Schröt.

  • ‡‡

    Identification tentative.

  • §§

    Porphyrellus porphyrosporus (Fr.) Gilb.

Albatrellus avellaneus PouzarEcMHoh6 4.3 ± 1.1−25.1 ± 0.8
Amanita constricta Thiers & AmmiratiEcMHoh5 1.8 ± 1.3−25.5 ± 0.5
Amanita franchetii (Boud.) FayodEcMHoh5 1.1 ± 0.9−25.4 ± 0.6
Amanita fulva (Schaeff.) Pers.EcMHoh5 1.5 ± 1.9−25.6 ± 0.7
Amanita pachycolea DE Stuntz in Thiers & AmmiratiEcMHoh6 1.5 ± 1.9−25.8 ± 1.3
Armillaria nabsnona TJ Volk & Burds.SapWHoh2 0.6 ± 1.2−23.2 ± 0.3
Armillaria ostoyae (Romagn.) HerinkSapW/ParHoh4−0.6 ± 1.1−22.2 ± 0.9
Boletopsis leucomelaena (Pers.) FayodEcMDP412.3 ± 2.0−25.5 ± 0.7
Boletus chrysenteron Bull.§EcMHoh4 4.9 ± 0.2−25.6 ± 0.6
Boletus coniferarum EA Dick & SnellEcMHoh311.0 ± 0.2−25.3 ± 0.1
Boletus edulis Bull. Fr.EcMHoh1 3.8−25.9
Boletus mirabilis MurrillEcMDP5 3.7 ± 0.8−26.4 ± 0.3
Boletus truncatus (Singer, Snell & EA Dick in Snell) PouzarEcMHoh2 9.4 ± 5.3−25.0 ± 0.2
Boletus zelleri Murrill**EcMHoh5 4.9 ± 1.1−24.6 ± 0.5
Bondarzewia mesenterica (Schaeff.) Kreisel?Hoh1 4.5−22.3
Cantharellus formosus CornerEcMDP5 5.0 ± 1.8−26.5 ± 0.7
Cantharellus formosus CornerEcMHoh10 3.2 ± 1.7−25.8 ± 0.7
Chroogomphus tomentosus (Murrill) OK MillerEcMDP10−4.4 ± 1.0−26.1 ± 0.7
Chroogomphus tomentosus (Murrill) OK MillerEcMHoh3−0.7 ± 2.5−26.0 ± 0.3
Chrysomphalina aurantiaca (Peck) RedheadSapLHoh2−1.2 ± 1.1−23.7 ± 0.5
Clavulina cristata (Holmsk.: Fr.) J. Schröt.††EcMHoh10 1.6 ± 0.4−25.5 ± 0.6
Clitocybe subditopoda PeckSapHHoh3 0.5 ± 0.1−23.5 ± 0.2
Cortinarius acutus (Pers.: Fr.) Fr.EcMDP5 4.0 ± 0.8−25.5 ± 0.8
Cortinarius brunneus (Pers.: Fr.) Fr.EcMDP5 5.6 ± 1.2−25.3 ± 0.8
Cortinarius cacao-color AH SmithEcMDP1 5.9−25.0
Cortinarius cacao-color AH SmithEcMHoh3 6.7 ± 0.9−25.0 ± 0.7
Cortinarius calochrous (Pers.: Fr.) Fr.EcMDP2 5.3 ± 0.6−24.8 ± 0.9
Cortinarius calopus P Karst.EcMDP2 5.8 ± 0.2−25.0 ± 1.5
Cortinarius clandestinus KauffmanEcMDP5 8.1 ± 0.5−24.6 ± 0.5
Cortinarius gentilis (Fr.) Fr.EcMDP5 5.3 ± 0.5−26.6 ± 0.9
Cortinarius gentilis (Fr.) Fr.EcMHoh3 4.4 ± 1.3−25.1 ± 0.6
Cortinarius laniger Fr.EcMDP19 8.2 ± 0.9−25.4 ± 1.3
Cortinarius malachius (Fr.: Fr.) Fr.EcMHoh3 4.9 ± 3.1−25.5 ± 0.5
Cortinarius montanus KauffmanEcMDP3 6.3 ± 0.8−26.5 ± 1.7
Cortinarius montanus KauffmanEcMHoh4 5.0 ± 0.4−26.2 ± 0.1
Cortinarius muscigenus PeckEcMDP10 6.7 ± 0.9−26.6 ± 0.5
Cortinarius obtusus (Fr.) Fr.EcMDP2 4.9 ± 2.0−26.5 ± 0.3
Cortinarius obtusus (Fr.) Fr.EcMHoh3 6.9 ± 1.1−25.9 ± 0.2
Cortinarius olympianus AH SmithEcMDP5 6.0 ± 0.8−24.7 ± 0.5
Cortinarius saturninus (Fr.) Fr.EcMDP2 4.1 ± 1.7−25.4 ± 0.9
Cortinarius semisanguineus (Fr.) GilletEcMDP410.4 ± 8.8−26.3 ± 1.1
Cortinarius spadiceus Batsch: Fr.EcMHoh2 5.7 ± 0.3−24.5 ± 0.7
Cortinarius talus Fr.EcMDP2 7.0 ± 2.1−25.9 ± 1.3
Cortinarius vanduzerensis AH Smith & TrappeEcMHoh5 8.9 ± 1.8−24.7 ± 0.7
Cortinarius variosimilis MM Moser & AmmiratiEcMDP2−1.5 ± 0.1−23.1 ± 0.8
Cortinarius vibratilis (Fr.) Fr.EcMDP5 6.1 ± 2.7−26.1 ± 1.1
Craterellus tubaeformis (Bull.: Fr.) Quél.EcMDP5 5.0 ± 2.5−25.7 ± 1.3
Cystoderma amianthinum (Scop.) Konrad & Maubl.SapLDP3−2.3 ± 0.7−26.2 ± 0.5
Cystoderma amianthinum (Scop.) Konrad & Maubl.SapLHoh1−2.8−24.3
Cystoderma granulosum (Batsch: Fr.) KühnerSapLDP1−0.3−25.2
Cystoderma granulosum (Batsch: Fr.) KühnerSapLHoh1−2.1−24.2
Entoloma nitidum Quél.?Hoh3 3.9 ± 1.4−24.6 ± 2.6
Fomitopsis pinicola (Swartz: Fr.) P Karst.SapWHoh2−4.1 ± 0.7−22.8 ± 0.1
Ganoderma applanatum (Pers.) Pat.SapWHoh2−4.2 ± 1.4−22.8 ± 0.1
Ganoderma oregonense MurrillSapWHoh4−2.0 ± 0.7−21.2 ± 0.9
Gomphidius subroseus KauffmanEcMDP5 1.6 ± 1.2−26.7 ± 0.5
Gomphus clavatus (Pers.: Fr.) SF GrayEcMDP3 7.6 ± 0.2−24.1 ± 0.5
Gymnopilus penetrans (Fr.) MurrillSapWHoh3−1.5 ± 1.8−23.6 ± 1.3
Gymnopus acervatus (Fr.) MurrillSapWDP8 0.1 ± 1.5−22.8 ± 0.8
Gymnopus acervatus (Fr.) MurrillSapWHoh3 1.0 ± 0.7−22.2 ± 0.4
Hebeloma mesophaeum (Pers.: Fr.) Quél.EcMDP4 4.6 ± 2.5−24.6 ± 0.5
Hebeloma incarnatulum AH SmithEcMDP2 1.8 ± 1.2−24.6 ± 0.2
Heterobasidion annosum (Fr.) Bref.SapW/ParDP2−1.6 ± 0.7−24.5 ± 1.0
Heterobasidion annosum (Fr.) Bref.SapW/ParHoh1−4.5−25.1
Hydnellum aurantiacum (Alb. & Schwein.) P Karst.EcMDP5 7.6 ± 0.9−25.1 ± 0.8
Hydnellum caeruleum (Hornem.) P Karst.EcMDP5 8.4 ± 0.7−25.3 ± 0.9
Hydnellum peckii BankerEcMDP10 6.3 ± 1.0−25.5 ± 0.9
Hydnum umbilicatum PeckEcMHoh3 6.3 ± 1.7−25.0 ± 0.7
Hygrophoropsis aurantiaca (Wulfen: Fr.) J. Schröt.SapLHoh5−2.8 ± 1.2−22.0 ± 0.6
Hygrophorus bakerensis AH Smith & HeslerEcMDP5 3.5 ± 4.3−25.5 ± 0.6
Hygrophorus camarophyllus (Alb. & Schwein.: Fr.) Dumée, Grandjean & MaireEcMDP2 4.1 ± 3.0−25.5 ± 0.7
Hygrophorus chrysodon (Batsch: Fr.) Fr.EcMDP3 1.1 ± 4.0−25.7 ± 0.9
Hygrophorus eburneus (Bull.: Fr.) Fr.EcMDP5 3.1 ± 2.6−25.7 ± 0.7
Hygrophorus olivaceoalbus (Fr.: Fr.) Fr.EcMHoh5 2.7 ± 3.5−24.9 ± 1.3
Hygrophorus purpurascens (Alb. & Schwein.: Fr.) Fr.EcMDP2 7.6 ± 0.1−24.9 ± 0.1
Hypholoma capnoides (Fr.) P Kumm.SapWDP1−1.3−24.1
Hypholoma capnoides (Fr.) P Kumm.SapWHoh5−3.8 ± 0.4−21.9 ± 0.3
Hypholoma fasciculare (Huds.: Fr.) P Kumm.SapWDP1 0.1−21.4
Hypholoma fasciculare (Huds.: Fr.) P Kumm.SapWHoh5−0.5 ± 1.0−22.2 ± 0.6
Hypholoma marginatum (Pers.: Fr.) J. Schröt.SapLHoh5−2.8 ± 0.3−22.5 ± 0.5
Inocybe assimilata Britzelm.EcMHoh2 2.8 ± 0.4−24.7 ± 0.6
Inocybe geophylla (Sowerby: Fr.) P Kumm. var. lilacina (Peck) GilletEcMDP3 1.5 ± 2.7−24.5 ± 0.2
Inocybe hirsuta var. maxima AH SmithEcMHoh10 3.3 ± 1.9−24.6 ± 0.9
Inocybe olympiana AH SmithEcMHoh5 0.5 ± 1.4−24.9 ± 0.7
Inocybe rimosa (Bull.: Fr.) Quél.EcMHoh10 1.4 ± 1.6−24.7 ± 0.9
Inocybe vaccina KühnerEcMDP4 1.2 ± 2.1−24.6 ± 1.5
Inocybe xanthomelas Boursier & KühnerEcMHoh2 1.0 ± 0.9−26.2 ± 0.4
Inonotus tomentosus (Fr.) TengSapWDP3−2.1 ± 1.8−23.5 ± 0.7
Inonotus tomentosus (Fr.) TengSapWHoh1−3.9−21.3
Laccaria bicolor (Maire) PD OrtonEcMDP4 0.2 ± 1.2−25.1 ± 0.5
Laccaria bicolor (Maire) PD OrtonEcMHoh5−0.5 ± 1.0−25.3 ± 0.4
Lactarius fallax var. concolor AH Smith & HeslerEcMHoh5 1.1 ± 0.8−24.4 ± 1.0
Lactarius kauffmanii var. kauffmanii Hesler & AH SmithEcMDP5 5.3 ± 1.1−25.6 ± 0.6
Lactarius olivaceo-umbrinus Hesler & AH SmithEcMHoh5 1.8 ± 0.8−25.1 ± 0.5
Lactarius olympianus Hesler & AH SmithEcMDP10 5.3 ± 2.6−24.8 ± 0.8
Lactarius olympianus Hesler & AH SmithEcMHoh2 5.4 ± 0.4−24.9 ± 0.4
Lactarius pallescens var. pallescens Hesler & AH SmithEcMDP5 3.3 ± 2.1−25.1 ± 0.6
Lactarius pseudomucidus Hesler & AH SmithEcMDP5 2.9 ± 0.9−25.5 ± 1.1
Lactarius pseudomucidus Hesler & AH SmithEcMHoh3 1.5 ± 0.6−25.1 ± 0.8
Lactarius rubrilacteus Hesler & AH SmithEcMDP5 4.7 ± 1.3−24.3 ± 1.0
Lactarius subflammeus Hesler & AH SmithEcMHoh5 5.2 ± 0.9−25.8 ± 0.5
Laetiporus conifericola Burds. & BanikSapWHoh3−2.4 ± 2.4−21.8 ± 1.1
Lepiota magnispora MurrillSapHDP2−0.8 ± 0.3−23.4 ± 0.4
Lyophyllum connatum (Schumach.: Fr.) SingerSapHDP6 1.4 ± 1.8−20.9 ± 0.5
Lyophyllum semitale (Fr.) KühnerEcMDP511.7 ± 1.6−25.0 ± 1.1
Mycena amicta (Fr.) Quél.SapWHoh2−2.7 ± 0.8−22.8 ± 1.0
Mycena aurantiidisca MurrillSapLDP1−4.3−25.4
Mycena aurantiidisca MurrillSapLHoh1−3.4−24.9
Mycena clavicularis (Fr.) Gillet sensu AH SmithSapLDP4−4.0 ± 0.2−24.5 ± 0.4
Mycena clavicularis (Fr.) Gillet sensu AH SmithSapLHoh1−2.0−25.2
Mycena epipterygia (Scop.: Fr.) SF GraySapLHoh4−3.7 ± 0.4−22.8 ± 0.6
Mycena haematopus (Pers.: Fr.) P Kumm.SapWHoh4−2.9 ± 0.2−22.7 ± 0.4
Mycena maculata P Karst.SapWDP3−1.0 ± 0.1−22.1 ± 0.4
Mycena strobilinoides PeckSapLDP2−3.3 ± 0.4−23.2 ± 0.7
Naucoria escharoides (Fr.: Fr.) P Kumm.EcMHoh3 4.3 ± 0.7−25.7 ± 0.8
Oligoporus perdelicatus (Murrill) Gilb. & RyvardenSapWHoh1−3.5−20.4
Paxillus atrotomentosus (Batsch: Fr.) Fr.SapWDP2−1.3 ± 0.5−22.4 ± 0.8
Phaeocollybia attenuata (AH Smith) SingerEcMHoh6 9.1 ± 0.3−24.3 ± 0.2
Phaeocollybia benzokauffmanii NorvellEcMHoh112.8−23.5
Phaeocollybia fallax AH SmithEcMHoh3 6.9 ± 1.9−24.3 ± 0.2
Phaeocollybia gregaria AH Smith & Trappe‡‡EcMHoh113.0−23.8
Phaeocollybia kauffmanii (AH Smith) SingerEcMHoh211.0 ± 0.4−25.4 ± 0.4
Phaeocollybia lilacifolia AH SmithEcMHoh210.5 ± 0.1−26.0 ± 0.3
Phaeocollybia piceae AH Smith & TrappeEcMHoh3 7.0 ± 0.8−25.3 ± 0.5
Phaeolus schweinitzii (Fr.) Pat.SapWDP4−0.2 ± 1.0−23.0 ± 1.0
Phaeolus schweinitzii (Fr.) Pat.SapWHoh2 0.0 ± 1.4−22.9 ± 1.7
Pholiota astragalina (Fr.) SingerSapWHoh5−3.3 ± 1.0−22.6 ± 0.7
Pholiota decorata (Murrill) AH Smith & HeslerSapLDP5−2.0 ± 0.8−23.7 ± 0.5
Pholiota decorata (Murrill) AH Smith & HeslerSapLHoh5−2.6 ± 0.6−23.1 ± 1.0
Pholiota flammans (Batsch: Fr.) P Kumm.SapWHoh4−3.7 ± 0.8−22.7 ± 0.3
Pholiota flavida (Schaeff.: Fr.) SingerSapWHoh5−1.0 ± 1.1−22.0 ± 0.8
Pholiota spumosa (Fr.) SingerSapWHoh3−1.9 ± 0.8−21.4 ± 0.1
Phylloporus rhodoxanthus (Schwein.) Bres.?DP1 0.7−26.0
Phylloporus rhodoxanthus (Schwein.) Bres.?Hoh1−1.3−25.7
Pleurocybella porrigens (Pers.: Fr.) SingerSapWHoh5−1.1 ± 1.8−21.5 ± 0.7
Pluteus cervinus (Schaeff.) P Kumm.SapWDP1−1.8−22.6
Pluteus cervinus (Schaeff.) P Kumm.SapWHoh3−2.0 ± 1.2−23.8 ± 0.9
Polyporus melanopus Pers.: Fr.SapWDP1 0.1−23.8
Polyporus melanopus Pers.: Fr.SapWHoh1−2.9−25.1
Pseudoplectania melaena (Fr.) Sacc.SapWDP1 0.8−24.9
Pseudoplectania melaena (Fr.) Sacc.SapWHoh2−2.4 ± 0.1−25.4 ± 1.5
Ramaria testaceoflava (Bres.) Corner var. brunnea (Zeller) Marr & DE StuntzEcMDP8 6.0 ± 1.4−25.4 ± 0.7
Rhodocollybia extuberans (Fr.) LennoxSapHDP3 2.2 ± 0.6−21.8 ± 0.5
Rhodocollybia extuberans (Fr.) LennoxSapHHoh3−1.4 ± 0.4−23.6 ± 0.5
Rhodocollybia maculata (Alb. & Schwein.: Fr.) SingerSapWDP2−0.5 ± 1.8−21.0 ± 0.3
Russula adusta (Pers.: Fr.) Fr.EcMDP5 4.3 ± 0.5−26.8 ± 0.8
Russula aeruginea Lindblad in Fr.EcMDP2 2.2 ± 1.4−26.0 ± 0.1
Russula bicolor Burl.EcMHoh4−1.3 ± 1.8−26.7 ± 0.9
Russula brevipes PeckEcMDP20 3.5 ± 0.8−24.8 ± 0.9
Russula brevipes PeckEcMHoh3 6.0 ± 0.9−24.4 ± 0.5
Russula crassotunicata SingerEcMHoh3 0.4 ± 1.3−26.4 ± 1.2
Russula fragilis (Pers.: Fr.) Fr.EcMDP9 1.6 ± 0.6−26.0 ± 0.6
Russula fragilis (Pers.: Fr.) Fr.EcMHoh5−0.6 ± 0.9−25.5 ± 0.7
Russula laurocerasi MelzerEcMHoh10 4.7 ± 1.0−24.9 ± 0.9
Russula nigricans (Bull.: Fr.) Fr.EcMHoh8 4.6 ± 0.9−25.2 ± 0.8
Russula occidentalis SingerEcMDP3 0.5 ± 0.4−25.3 ± 0.7
Russula queletii Fr.EcMHoh2 1.0 ± 0.1−24.8 ± 0.0
Russula raoultii Quél.EcMHoh2−1.1 ± 0.1−27.3 ± 0.4
Russula xerampelina (Schaeff.) Fr.EcMDP5 4.0 ± 1.0−24.8 ± 0.5
Sarcodon fuscoindicum (KA Harrison) Maas Geest.EcMDP2 9.1 ± 1.5−25.3 ± 0.0
Sarcodon scabrosum (Secr.: Fr.) P Karst.EcMDP8 7.6 ± 0.7−24.9 ± 0.8
Suillus lakei (Murrill) AH Smith & ThiersEcMDP9 5.8 ± 1.5−24.5 ± 0.9
Suillus punctatipes (Snell & EA Dick) AH Smith & ThiersEcMDP1011.6 ± 1.1−25.0 ± 0.6
Tricholoma atroviolaceum AH SmithEcMDP1 7.5−27.3
Tricholoma atroviolaceum AH SmithEcMHoh211.0 ± 1.1−25.7 ± 0.1
Tricholoma flavovirens (Pers.: Fr.) S. LundellEcMDP2 7.7 ± 0.3−25.3 ± 1.0
Tricholoma focale (Fr.) RickenEcMDP5 9.7 ± 1.9−26.0 ± 0.8
Tricholoma inamoenum (Fr.: Fr.) GilletEcMDP5 5.4 ± 0.7−25.3 ± 0.7
Tricholoma magnivelare (Peck) RedheadEcMDP1112.1 ± 2.9−25.7 ± 0.5
Tricholoma myomyces (Pers.: Fr.) JE LangeEcMDP3 6.4 ± 3.0−25.5 ± 1.1
Tricholoma nigrum Shanks & OvreboEcMDP2 9.7 ± 1.4−26.3 ± 0.3
Tricholoma pardinum (Pers.) Quél.EcMDP2 7.6 ± 0.8−24.9 ± 0.2
Tricholoma saponaceum (Fr.: Fr.) P Kumm.EcMDP4 7.7 ± 1.9−24.2 ± 1.2
Tricholoma saponaceum (Fr.: Fr.) P Kumm.EcMHoh510.3 ± 3.2−25.3 ± 0.5
Tricholoma sejunctum (Sowerby: Fr.) Quél.EcMHoh5 6.8 ± 2.6−25.5 ± 0.6
Tricholoma vaccinum (Schaeff.: Fr.) P Kumm.19DP2 6.8 ± 0.4−25.3 ± 1.1
Tricholoma virgatum (Fr.: Fr.) P Kumm.EcMDP4 9.8 ± 2.0−25.7 ± 1.3
Tricholoma virgatum (Fr.: Fr.) P Kumm.EcMHoh1 8.4−23.9
Tricholomopsis decora (Fr.) SingerSapWDP2−1.9 ± 0.3−22.7 ± 0.5
Tricholomopsis decora (Fr.) SingerSapWHoh2−3.8 ± 1.0−22.9 ± 1.6
Tylopilus porphyrosporus (Fr.) AH Smith & Thiers§§EcMHoh2 3.9 ± 0.9−24.4 ± 0.1
Xeromphalina campanella (Batsch: Fr.) Kühner & MaireSapWHoh4−3.0 ± 0.5−21.7 ± 0.9
Xeromphalina cornui (Quél.) J FavreSapLDP5−3.5 ± 0.8−23.5 ± 0.8
Xeromphalina cornui (Quél.) J FavreSapLHoh3−4.8 ± 0.1−23.0 ± 0.0

Ancillary

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