Controls of isotopic patterns in saprotrophic and ectomycorrhizal fungi

Soil Biology & Biochemistry 48 (2012) 60e68

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Controls of isotopic patterns in saprotrophic and ectomycorrhizal fungi Erik A. Hobbie a, *, Fernando S. Sánchez b, Paul T. Rygiewicz b a b

Complex Systems Research Center, University of New Hampshire, Durham, NH 03824, USA US Environmental Protection Agency, National Health and Environmental Effects Research Laboratory, Western Ecology Division, 200 SW 35th Street, Corvallis, OR 97333, USA

a r t i c l e i n f o

a b s t r a c t

Article history: Received 9 October 2011 Received in revised form 17 January 2012 Accepted 18 January 2012 Available online 3 February 2012

Isotopes of nitrogen (d15N) and carbon (d13C) in ectomycorrhizal and saprotrophic fungi contain important information about ecological functioning, but the complexity of physiological and ecosystem processes contributing to fungal carbon and nitrogen dynamics has limited our ability to explain differences across taxa. Here, we measured d15N and d13C in needles, litter, soil, wood, fungal caps, and fungal stipes at numerous forested sites in Oregon, USA to determine how functional attributes and biochemical processes may influence isotopic values. Ectomycorrhizal fungi were classified by hydrophobicity of ectomycorrhizae and by patterns of hyphal exploration; saprotrophic fungi were classified into wood decay and litter decay fungi. For d15N, caps of hydrophobic taxa averaged 8.6&, hydrophilic taxa 3.2&, and saprotrophic taxa 0.5&, whereas needles averaged 3& and soil at 5e12 cm averaged 2&. Caps were higher in d15N, d13C, %N, and %C than stipes by 1.7&, 0.6&, 1.75%, and 2.61%, respectively, presumably because of greater protein content in caps than stipes. Isotopic enrichment of caps relative to stipes was greater in hydrophobic taxa (3.1& for 15N and 0.8& for 13C) than in hydrophilic taxa (1.1& for 15N and 0.5& for 13C). In multiple regressions, 45% of variance in d15Ncapestipe and 30% of variance in d13Ccapestipe was accounted for by various elemental, isotopic, and categorical variables. We estimated that fungal protein was enriched in 15N relative to fungal chitin by 15& in hydrophobic taxa and by 7& in hydrophilic taxa. Fungal protein was enriched in 13C by 4.2
0.5& relative to carbohydrates. Isotopic signatures of sources and isotopic fractionation during metabolic processing influence both isotopic patterns of sporocarps and the isotopic partitioning between caps and stipes; functional groups differed in processing of both nitrogen isotopes and carbon isotopes. Ó 2012 Elsevier Ltd. All rights reserved.

Keywords: Organic nitrogen use Sporocarps Protein Chitin Nitrogen isotopes Carbon isotopes Coniferous forests

1. Introduction Comparing isotopic patterns in fungal sporocarps to those of ecosystem components has provided insight into the roles of both ectomycorrhizal and saprotrophic fungi in ecosystem carbon and nitrogen cycling (Hobbie et al., 1999; Högberg et al., 1999a; Kohzu et al., 1999; Taylor and Fransson, 2006). Here, we use an extensive data set of isotopic ratios in soil, wood, foliage, and sporocarps of many taxa of ectomycorrhizal and saprotrophic fungi to assess the probable mechanisms controlling fungal d15N and d13C. We will assess how internal processes in sporocarps affect d15N and d13C by comparing isotopic patterns in caps and stipes to elemental patterns and life history strategies. Several factors control d15N values in fungi. The d15N of source nitrogen should influence fungal d15N, with bulk d15N increasing with depth in soil profiles (Hobbie and Ouimette, 2009). In * Corresponding author. E-mail address: [email protected] (E.A. Hobbie). 0038-0717/$ e see front matter Ó 2012 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2012.01.014

addition, the d15N of organic nitrogen is generally higher than inorganic nitrogen (Koba et al., 2003; Takebayashi et al., 2010), but varies with the relative fluxes of ammonification, nitrification, and denitrification (Houlton et al., 2007). For ectomycorrhizal fungi, the isotopic signature of the original nitrogen source can be modified by fractionation against 15N during transfer of nitrogen compounds to host plants (Hobbie et al., 2000). Relative to available nitrogen sources, ectomycorrhizal fungi will then be enriched in 15N and their ectomycorrhizal hosts will be depleted in 15N. However, even in saprotrophic fungi, sporocarps are enriched at least 3& in 15N relative to bulk soil nitrogen, despite not transferring 15N-depleted nitrogen to plants (Hobbie et al., 2005). Internal 15N fractionation may contribute to this enrichment if fungi partition nitrogen into 15 N-depleted chitin and 15N-enriched protein (Taylor et al., 1997) and the protein is preferentially mobilized for sporocarp formation. The growth patterns of ectomycorrhizal fungi also appear to influence d15N of sporocarps. Agerer (2001, 2006) classified ectomycorrhizal fungi based on whether mycorrhizae were hydrophobic or hydrophilic and the extent and form of hyphal development

E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

(termed exploration type). This morphological classification corresponds to functionality, with the hydrophobic exploration types (medium-distance fringe, medium-distance mat, and long-distance) putatively possessing better proteolytic capabilities than hydrophilic exploration types (contact, short-distance, and medium-distance smooth) (Lilleskov et al., 2011). Nitrogen isotope patterns in ectomycorrhizal sporocarps correspond with these exploration types, with hydrophobic exploration types generally 4e7& higher in d15N than hydrophilic exploration types (Hobbie and Agerer, 2010). Differences in isotopic ratios between caps and stipes of sporocarps could potentially provide additional insight. These components differ in their chemical constituents, nitrogen concentration, and d15N. In four ectomycorrhizal species, Taylor et al. (1997) attributed an average 15N enrichment of caps relative to stipes of 2.2
0.2& to a concurrently measured 15N enrichment of protein relative to chitin of 9.9
0.5& and a greater proportion of protein in caps than in stipes. Because only three species (all ectomycorrhizal) have been tested to date, these 15N enrichments of caps relative to stipes have not been compared across different functional classifications, such as between saprotrophic and ectomycorrhizal fungi, or among ectomycorrhizal fungi of different exploration type. 13 C enrichment patterns in caps and stipes have yet to be measured, but 13C enrichment in protein relative to chitin in other organisms such as locusts (Webb et al., 1998), suggests that caps should also be enriched in 13C relative to stipes. Relative to stipes, caps appear higher in protein and lipids and lower in fiber (Alam et al., 2008). The mechanism of N-acetyl glucosamine synthesis (Carlile et al., 2001) suggest that fractionation against 15N during aminotransferase of nitrogen from glutamine to N-acetyl glucosamine accounts for the consistent depletion of 15N in chitin relative to protein, muscle, or bulk tissue in diverse heterotrophic organisms (Macko et al., 1986; Webb et al., 1998; Hobbie and Colpaert, 2003). No CeN bond is involved in chitin synthesis from N-acetyl glucosamine, so the 15N depletion of chitin must arise during N-acetyl glucosamine formation. Similarly, 13C enrichment of amino acids relative to chitin and other carbohydrates during metabolic processing drives the 13 C depletion of these carbohydrates relative to protein and muscle in heterotrophic organisms (Schimmelmann and DeNiro, 1986; Webb et al., 1998; Schimmelmann, 2011). According to the above arguments, %N and d15N of caps will differ from stipes if the relative proportion of chitin and protein in these two fungal components also differ, even if the 15N difference between protein and chitin is held constant. However, the d15N of the two compound classes (protein and chitin) can also vary (Taylor et al., 1997). For a given relative difference in %N between caps and stipes, an increased difference in d15N indicates increased isotopic separation between chitin and protein, if the relative increase in protein is similar. How isotopes partition between two components (such as chitin and protein) depends on whether the system is treated as open or closed. For an open system, isotopic differences between the two components will remain constant (Fig. 1a), whereas isotopic differences for the closed system will vary depending on the relative allocation between the two components (Fig. 1b). Closedsystem mathematics have been used to explain d15N patterns in sporocarps (Hobbie et al., 2005), in which increased sequestration of 15N-depleted chitin in fungal hyphae increases the 15N enrichment of sporocarps relative to available nitrogen sources (Trudell et al., 2004; Hobbie and Agerer, 2010). We suggest that the extensive hyphal development in hydrophobic systems will lead to increased chitin sequestration and consequently increased 15N and 13 C enrichment of fungal protein relative to hydrophilic systems, and can be treated as a closed system. This isotopic partitioning will accordingly increase the isotopic enrichment of caps relative to stipes.

61

Fig. 1. Theoretical relationship between nitrogen isotope ratios in protein and chitin for open versus closed systems as the proportion of system nitrogen as chitin increases from 0 to 1 (expresses as the transfer ratio, Tr). This assumes that fractionation against 15N is 10& during synthesis of chitin from protein. a. Open system, d15Nproteinechitin ¼ Dchitin. b. Closed system, d15Nprotein-accumulated chitin ¼ Dchitin/Tr$ln [1/(1 e Tr)].

Mathematical formulations for isotopic patterns in open and closed systems are given in Fry (2006). For an open system in which some nitrogen from protein can be transformed into chitin with a fractionation against 15N during chitin synthesis of Dchitin, the following equations apply, in which Tr is the proportion of reactant transformed into product:

d15 Nchitin ¼ d15 Nsource N  ð1  Tr Þ$Dchitin

(1)

d15 Nprotein ¼ d15 Nchitin þ Dchitin

(2)

15

Thus, the N enrichment of protein relative to chitin is a constant, equivalent to Dchitin. For a closed system, the 15N enrichment of protein relative to chitin is not a constant, although the 15N depletion during chitin synthesis remains the same.

d15 Nprotein ¼ d15 Nsource N  Dchitin $lnð1  Tr Þ

(3)

d15 Naccumulated chitin ¼ d15 Nsource N þ Dchitin $ð1=Tr  1Þ$½lnð1  Tr Þ

(4)

d15 Nproteinaccumulated chitin ¼ Dchitin =Tr $ln½1=ð1  Tr Þ

(5)

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E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

In this study, we collected sporocarps together with other ecosystem components at 30 sites in Oregon, USA. Sites ranged from humid coastal forests to the drier interior forests east of the Cascade Mountains (Bowling et al., 2002). We compared the d15N and d13C of ectomycorrhizal and saprotrophic sporocarps with foliage, litter, wood, and soil. We assessed differences in %N, d15N, %C, and d13C between caps and stipes for insight into mechanisms creating isotopic patterns in different fungal taxa. Based on the above arguments, we propose the following hypotheses: 15 N enrichment in caps versus stipes corresponds to the relative increase in %N between caps and stipes. This arises because increased %N is caused by increased protein, and protein is high in d15N. (2) Because protein is also high in d13C, caps will be higher in d13C than stipes. (3) Isotopic patterns during fungal metabolism can be treated assuming a closed system during sporocarp formation, and assuming greater chitin sequestration in hydrophobic taxa than in hydrophilic taxa. Accordingly, 15N and 13C enrichment in caps versus stipes will be higher in hydrophobic taxa than in hydrophilic taxa.

(1)

2. Methods 2.1. Sample collection At 30 different locations across Oregon during 1998 and 1999, sporocarps of 193 taxa of ectomycorrhizal fungi, litter decay fungi, and wood decay fungi were collected, totalling 337 specimens. For 21 of the collections, exact location was not recorded. Most collecting sites had forests dominated by Pseudotsuga menziesii (Mirb.) Franco (Douglas-fir), Tsuga heterophylla (Raf.) Sarg. (western hemlock), Picea sitchensis (Bong.) Carr. (Sitka spruce), or Pinus ponderosa C. Lawson (Ponderosa pine). Collections were identified, cleaned of litter and humus, and freeze-dried. One specimen of each collection was divided into cap and stem and ground using a mortar and pestle. Ectomycorrhizal species were classified by exploration type and hydrophobicity according to Agerer (2001, 2006) and Hobbie and Agerer (2010). Species of unknown exploration type were assigned to the dominant exploration type in the genus. The genera for the three hydrophilic exploration types were (1) contact: Chroogomphus, Hygrophorus, Lactarius; (2) short-distance: Inocybe, Rozites; (3) medium-distance smooth: Albatrellus, Amanita, Cantharellus, Gomphidius, Laccaria, Russula. The genera for the three hydrophobic exploration types were (4) medium-distance fringe: Cortinarius, Hebeloma, Hydnum, Tricholoma; (5) medium-distance mat: Gomphus, Ramaria, Sarcodon; and (6) long-distance: Boletus, Leccinum, Paxillus, Suillus. Because of uncertain species identification, all Entoloma samples were excluded from analysis. The pick-a-back exploration type classification for Chroogomphus and Gomphidius was not used, since assigning fungi to this type requires examination of root tips. At 21 sites (17 of which were also sampled for fungi), soil samples (five per site, approximately 200 g fresh weight) were collected at two depths, 0e5 cm and 5e12 cm. Five samples of green mature needles were also collected at each site from the bottom 5 m of canopy. Most forests sampled were dominated by a single tree species, but collections were from a range of species in mixed forests. Five samples of surface litter (mixed foliar and twig litter) were also taken from each site (100 g dry weight). We collected five samples per site (100 g dry weight) of downed, decayed wood. Samples were dried at 60  C for 48 h, and then ground using a rotating sample mill.

2.2. Sample analysis Samples were analyzed for d15N, d13C, %N, and %C on a Finnigan Delta-Plus isotope ratio mass spectrometer linked to a Carlo Erba NC2500 elemental analyzer (Finnigan MAT GmbH, Bremen, Germany) located at the U.S. EPA, Corvallis, Oregon, USA. The internal standards for isotopic and concentration measurements were acetanilide and spinach (NIST 1570a). We report stable isotope abundances as d15N (or d13C) ¼ (Rsample/Rstandard  1)$1000, where R ¼ 15N/14N or 13C/12C of either the sample or the reference standard (atmospheric N2 for nitrogen, PeeDee belemnite for carbon). The precision of isotopic measurements based on duplicate samples was
0.2& for 15N and 13C. When comparing between samples, samples with more of the heavy isotope are referred to as heavier, or enriched; samples with more of the light isotope are lighter, or depleted. If isotopic fractionation (D) is calculated between two pools (such as between protein and chitin), it is calculated as D ¼ (dsubstrate  dproduct)/(1 þ dsubstrate/1000&). Nitrogen concentrations in wood were too low for adequate %N and d15N analyses. To compare the relative difference in nitrogen between caps and stipes across taxa, the concentration difference was calculated as (1  %Nstipe/%Ncap). Most data analyses used the statistical package Statview (Abacus Concepts, Berkeley, California). Elemental and isotopic differences between caps and stipes were also calculated. Differences in isotopic values and elemental concentrations were evaluated using one-way ANOVAs or t tests if only two groups were compared, and variability within groups was compared using an F-test. Means among classified groups were compared using a post hoc TukeyeKramer test at the 0.05 significance level. Relationships between variables were compared using correlations. Stepwise, forward linear multiple regressions only used data from known locations (the 21 sporocarps of unknown location were excluded), and explained variability in d15Ncapestipe and d13Ccapestipe using the SigmaPlot program (Systat, San Jose, California). Both regression models used the following continuous variables: %Ncapestipe, 1  %Ncap/%Nstipe, and %Ccapestipe. The model to explain d15Ncapestipe included d13Ccapestipe as an independent variable and the model to explain d13Ccapestipe included d15Ncapestipe as an independent variable. To determine which independent variables to retain in the multiple regression, we used the SigmaPlot defaults of setting the “F-to-remove” at a value of 4.0, and the “F-to-Enter” at 3.9. Three categorical variables were also included in the multiple regressions, corresponding to the main classifications of the fungi, such as having hydrophilic or hydrophobic ectomycorrhizae, or being a litter decay fungus, with a value of 1 assigned to the appropriate categorical variable and other categorical variables assigned a value of 0. Wood decay fungi were assigned a value of 0 for all three categorical variables. 2.3. Theoretical relationship between nitrogen isotope composition in caps and stipes We sought to relate cap and stipe isotopic composition to the isotopic composition of protein and chitin and stipe and cap %N. Nucleic acids appear to be less than 10% (by mass) of protein content in gills of Coprinus lagopus (Iten and Matile, 1970; they recorded maximum concentrations of 10%, 2%, and 0.8% for protein, N-acetyl glucosamine, and RNA in cell-free extracts). Because gills are the site of fungal reproduction, nucleic acid content should be higher in gills than in other tissues. For this analysis, we assume that nitrogen in fungi is composed of two main pools, chitin and protein, and assume that stipe nitrogen consists of a fungal chitin pool (a, stipechitin) and a fungal protein pool (b, stipeprotein). Cap nitrogen retains these two pools but adds additional protein (c, capprotein). We also assume that d15N values of protein and chitin do not change

E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

between caps and stipes, with protein 10& enriched in 15N relative to chitin. Under these assumptions, stipe and cap d15N can be expressed as:

d15 Nstipe ¼ ½b=ða þ bÞ$ 10% þ d15 Nchitin

(6)

d15 Ncap ¼ ½ðb þ cÞ=ða þ b þ cÞ$ 10% þ d15 Nchitin

(7) 15

The difference between these two measurements, d Ncap 

d15Nstipe, can be expressed as:

d15 Ncap  d15 Nstipe ¼ a=ða þ bÞ$c=ða þ b þ cÞ$10%

(8)

%Nstipe equals 100%$(a þ b)/total biomass and %Ncap equals 100%$(a þ b þ c)/total biomass. Accordingly, the quantity, c/(a þ b þ c), is equivalent to 1  %Nstipe/%Ncap.




d15 Ncap  d15 Nstipe ¼ a=ða þ bÞ$ 1  %Nstipe =%Ncap $10%

(9)

This can also be expressed as:



d15 Ncap  d15 Nstipe ¼ stipechitin = stipechitin þ stipeprotein
$ 1  %Nstipe =%Ncap $10%

 ð9aÞ

In this analysis, the isotopic difference between caps and stipes can be reduced to three factors: 1  %Nstipe/%Ncap, the proportion of stipe nitrogen that is chitin, and the 15N enrichment of protein relative to chitin. 3. Results 3.1. Plant litter and soil Plant litter, wood, and soil were collected at 21 sites. Concentrations of nitrogen and carbon decreased from litter (0.99
0.24 % N, 44.39
4.52 %C) to 0e5 cm soil (0.69
0.44%, 17.00
11.78 %C) to 5e12 cm soil (0.29
0.27 %N, 7.20
7.86 %C). d13C values increased from needles to litter to soil to wood, with overall averages of 28.8
1.7& for needles, 27.1
0.9& for litter, 26.3
0.9& for 0e5 cm soil, 25.5
0.8& for 5e12 cm soil, and 24.8
1.1& for wood (all values
SD). d15N values increased from needles to litter to soil, with overall averages of 3.3
1.3& for needles, 2.3
0.9& for litter, 0.6
1.2& for 0e5 cm soil, and 2.6
1.2& for 5e12 cm soil. Elemental and isotopic data for the 21 sites are given in Appendix 1.

63

litter decay (22.8
0.3&) to ectomycorrhizal fungi (24.7
0.1&) (Fig. 2). In ectomycorrhizal fungi, caps of hydrophobic taxa averaged 8.5& for d15N and 24.6
0.1& for d13C, whereas caps of hydrophilic taxa averaged 3.2
0.2& for d15N and 24.8
0.1& for d13C. Caps averaged higher than stipes by 1.7& in d15N, 0.6& in d13C, 1.75% in %N, and 2.61% in %C. The isotopic enrichment of caps relative to stipes (designated d15Ncapestipe and d13Ccapestipe) was higher in ectomycorrhizal fungi than in litter decay or wood decay fungi, and in ectomycorrhizal fungi, was significantly greater in hydrophobic exploration types (3.1& in d15N and 0.8& in d13C, p < 0.001 for both) than in hydrophilic exploration types (1.1& in d15N and 0.5& in d13C) (Fig. 2). d15Ncapestipe varied within the different exploration types, ranging from 1.0& for medium-distance smooth to 3.6& for medium-distance fringe. d13Ccapestipe also varied by exploration type, ranging from 0.3& for contact exploration types to 0.9& for medium-distance fringe exploration types (Table 1). The relative differences in nitrogen between caps and stipes was calculated as (1  %Nstipe/%Ncap) and varied by exploration type in ectomycorrhizal fungi. It ranged from 0.29 for short-distance to 0.45 for medium-fringe exploration types, with litter decay fungi lower (0.36) than wood decay fungi (0.42). This quantity was significantly higher in fungi with hydrophobic ectomycorrhizae (0.436
0.020) than in fungi with hydrophilic ectomycorrhizae (0.351
0.012) (t-test, df ¼ 139, p < 0.001) (Table 1). d15Ncapestipe and d13Ccapestipe were positively correlated in ectomycorrhizal fungi (r2 ¼ 0.16, n ¼ 210, p < 0.001) and wood decay fungi (r2 ¼ 0.38, n ¼ 39, p < 0.001) but were uncorrelated in litter decay fungi (r2 ¼ 0.01, n ¼ 59, p ¼ 0.451). The relationship between d15Ncapestipe and d13Ccapestipe in different genera is shown by nutritional source, hydrophobicity, and exploration type in Fig. 3. We used multiple regressions to explore the potential controls and covariates of the isotopic difference between caps and stipes for both nitrogen and carbon (d15Ncapestipe and d13Ccapestipe). The multiple regressions explaining the maximum amount of variance are shown in Table 2. Only the statistically significant variables have been retained in the table. For d15Ncapestipe, the significant factors (in descending order of importance) were the relative proportion of nitrogen in caps versus stipes, %Ncapestipe, d13Ccapestipe, the hydrophobicity of ectomycorrhizae, %Ccapestipe, and the hydrophilicity of

3.2. Fungi A total of 337 collections of fungi were made at 27 sites, of which 227 were ectomycorrhizal, 63 were litter decay, and 47 were wood decay fungi. Ectomycorrhizal fungi were classified by hydrophobicity of ectomycorrhizae and by exploration type (Hobbie and Agerer, 2010), with 157 specimens classified as having hydrophilic ectomycorrhizae and 70 as having hydrophobic ectomycorrhizae. The most common exploration type was medium-distance smooth, of which 98 were collected, followed in abundance by medium-distance fringe (43), contact (33), short-distance (26), long-distance (23), and medium-distance mat (4). Complete details on locations sampled and species collected are given in Appendix 2. Nitrogen and carbon isotopic signatures and concentrations varied across functional class, exploration type, and fungal tissue. Caps averaged 4.49
1.57% (SD) in %N and 41.16
2.75 (SD) in %C, whereas stipes averaged 2.83
1.19% (SD) in %N and 38.57
2.36 (SD) in %C. In caps of sporocarps, d15N increased from wood decay (0.4
0.4&) to litter decay (1.3
0.5&) to ectomycorrhizal fungi (4.9
0.3&) and d13C decreased from wood decay (22.4
0.3&) to

Fig. 2. d13C and d15N values of different ecosystem pools (needles, litter, wood, 0e5 cm soil, and 5e12 cm soil) and of caps and stipes of different fungal groups. Fungi are separated into wood decay fungi (wood d), litter decay fungi (litt d), fungi with hydrophobic ectomycorrhizae (ECM-ho), and fungi with hydrophilic ectomycorrhizae (ECM-hi). Names of genera are indicated on the Figure. Wood d15N was not measured; it is indicated by a box spanning its probable range between the d15N of needles and the d15N of caps of wood decay fungi.

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E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

Table 1 d15N and d13C in caps, the enrichment in caps versus stipes for 15N, 13C, %N, and %C, and the relative enrichment of caps versus stipes for nitrogen concentrations. Because of missing variables during measurement or, in some fungi, the absence of stipes, the number of measurements varies for different variables. Number of samples (n) is given in parentheses for each parameter in the table, number of samples for each fungal type is given in the second column. Ectomycorrhizal fungi are classified by exploration type, saprotrophic fungi by substrate (litter or wood). Significance values are given in italics for ANOVAs within exploration types for ectomycorrhizal fungi or between the two saprotrophic decay types (unpaired t-test); significance values are bolded if p < 0.05. Significant differences among exploration types or between the two saprotrophic types are indicated by different superscripted letters (a, b, and c), with no letter indicating no significant differences. Exploration or Decay Type n Ectomycorrhizal contact medium-smooth short medium-fringe medium-mat long Saprotrophic litter decay wood decay

n

33 98 26 43 4 23 63 45

d15Ncap (&) (333)

d13Ccap (&) (336)

d15Ncapesti (&) (309)

d13Ccapesti (&) (335)

%Ncapesti (%) (312)

%Ccapesti (%) (334)

1  %Nsti/cap (312)

<0.001 2.8
0.6a 3.5
0.3a 2.4
0.4a 8.5
0.5b 13.5
1.3c 7.8
0.6b 0.022 1.3
0.5b 0.4
0.3a

0.002 25.0 24.8 24.0 24.8 24.3 24.3 0.334 22.8 22.4

<0.001 1.4
0.2b 1.0
0.1a 1.5
0.2ba 3.6
0.3c 3.6
0.1cb 2.3
0.3b 0.022 1.4
0.2 1.9
0.2

<0.001 0.3
0.1a 0.5
0.1a 0.6
0.1 0.9
0.1b 0.9
0.2 0.7
0.2 0.968 0.7
0.1 0.7
0.1

<0.001 1.8
0.1 1.4
0.1a 1.4
0.1a 1.7
0.1 1.7
0.2 2.2
0.2b 0.178 2.1
0.1 1.8
0.2

0.036 2.9
0.2 3.2
0.2b 2.9
0.4 1.9
0.8 3.0
0.9 1.6
0.3a 0.284 2.5
0.4 1.6
0.4

<0.001 0.42
0.02bc 0.35
0.01ab 0.29
0.02a 0.45
0.02c 0.43
0.05 0.40
0.03bc 0.045 0.37
0.01a 0.42
0.03b









0.3a 0.1 0.3b 0.2 0.3b 0.3b


0.3
0.3

ectomycorrhizae. For d13Ccapestipe, the significant factors were % Ncap-stipe, d15Ncapestipe, and %Ccapestipe. Adjusted r2 versus the independent variables is 0.452 for d15Ncapestipe and 0.306 for d13Ccapestipe (n ¼ 286, p < 0.001 for both). Intercepts for both multiple regressions were not significantly different from zero. 4. Discussion 4.1. Ecosystem patterns in d13C Within a site, increases among ecosystem pools in d13C going from needles to litter to deeper soil horizons presumably reflect the transport of 13C-enriched carbohydrates within plants from leaves to roots and wood (Hobbie and Werner, 2004) and the routing of root-derived carbon into deeper soil horizons via mycorrhizal fungi and root decay. In addition, the retention of 13C-enriched carbon derived from microbes and microbial protein will increase d13C values of soil as material is increasingly processed by microbes

(Boström et al., 2007; Sollins et al., 2009). Similar d13C values for ectomycorrhizal fungi and wood may reflect the common carbon source (tree-transported sugars) and similar levels of fractionation during metabolism by trees during wood formation and by fungi during sporocarp formation, despite the rather different chemical constituents of these two pools (cellulose and lignin for trees, carbohydrates and protein for fungi). d13C values for needles are probably somewhat low relative to other ecosystem pools because we were unable to sample overstory foliage, which is generally 13 C-enriched by 1& or more relative to mid-story and understory foliage (Gottlicher et al., 2006). The higher d13C values of litter and wood decay fungi than ectomycorrhizal fungi presumably reflect the incorporation of carbon into saprotrophic biomass from 13C-enriched wood cellulose rather than from 13C-depleted wood lignin and lipids (Gleixner et al., 1993; Hobbie, 2005). Tree-transported sugars assimilated by ectomycorrhizal fungi are probably 13C-depleted relative to those used to make wood cellulose because a substantial portion of the sugars used for wood are diverted to make 13C-depleted secondary compounds in wood, resulting in 13C enrichment of the remaining sugars used to make wood cellulose (Hobbie and Werner, 2004). These explanations are shown schematically in Fig. 4, with the small difference in 13C enrichment between hydrophobic and hydrophilic taxa explained in Section 4.5. The similar d13C values for litter decay and wood decay fungi reflect the common carbon source, cellulose, for both these fungal types. Although division of saprotrophic fungi into only two types simplifies the continuum of decay types and nutrient sources in these fungi, this approach is nonetheless useful as a classification tool. 4.2. Ecosystem patterns in d15N

Fig. 3. Isotopic differences in carbon and nitrogen between caps and stipes, plotted by genus for n > 1. Mean values for the species  location combinations within a genus are plotted, standard errors are plotted in grey in the positive direction only for clarity. Black symbols represent taxa with hydrophobic mycorrhizae, for medium-distance fringe taxa, squares (-); medium-distance mat, triangles (:); and long-distance, circles (C). Clear symbols represent taxa with hydrophilic mycorrhizae, for mediumdistance smooth taxa, squares (,); short-distance, triangles (D); and contact, circles (B). Litter decay taxa are indicated with a gray x, wood decay fungi with a grey cross. Representative genera are named on the graph. Complete data are given in Appendix 2.

Progressive 15N enrichment from needles to litter to deeper soil horizons (Fig. 2) reflects repeated transfer of 15N-depleted nitrogen from ectomycorrhizal fungi to trees and 15N enrichment during microbial processes (Hobbie and Ouimette, 2009). The resulting 15 N-depleted litter then accumulates at the soil surface and 15 N-enriched fungal residues accumulate in deeper soil horizons (Högberg et al., 1996; Wallander et al., 2009). The higher d15N of saprotrophic fungi growing on humus and litter compared to wood decay fungi has been reported previously (Gebauer and Taylor, 1999; Kohzu et al., 1999; Taylor et al., 2003; Trudell et al., 2004). This pattern could reflect differences in the nitrogen sources used by these fungal types or in the relative proportions of 15N-enriched protein and 15N-depleted chitin in

E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

65

Table 2 Stepwise multiple regressions to explore controls over d15Ncapestipe and d13Ccapestipe (n ¼ 286). The model explained 0.452 of the variance for d15Ncapestipe and 0.306 of the variance for d13Ccapestipe (adjusted r2). Dr2 is the increase in the % variance explained by adding the specified independent variable in the stepwise regression. Only statistically significant variables have been retained in the table. Independent variable

d15Ncapestipe Coefficient
SE

Intercept %Ncapestipe %Ccapestipe 1-%Nstipe/%Ncap d13Ccapestipe d15Ncapestipe Hydrophobic Hydrophilic a

0.300 0.781 0.123 5.171 0.853 e 1.084 0.571

d13Ccapestipe Coefficient Dr








0.242 0.126 0.034 0.706 0.149


0.207
0.175

2

e 4.9% 1.7% 11.0% 4.6% e 22.0% 2.2%

p


SE

0.215a <0.001 <0.001 <0.001 <0.001 e <0.001 <0.001

0.073 0.253 0.034 e e 0.121 e e


0.069
0.033
0.012


0.017

Dr2

p

e 17.5% 1.9% e e 11.9% e e

0.289a <0.001 0.006 e e <0.001 e e

p-values for intercepts are taken from non-stepwise multiple regression analyses.

different taxa. For example, in this study litter decay fungi were 1.3& and 2.1% higher in 15N and %N than wood decay fungi, and Taylor and Fransson (2006) reported quite similar enrichment patterns (1.8& and 2.5%). Thus, we cannot rule out that source differences in d15N between the two saprotrophic types may be small, and sporocarp d15N primarily reflects relative protein content, which will correlate with %N. However, %N and d15N of litter decay fungi were not highly correlated and the higher variability of litter fungi than wood decay fungi for these two parameters (F-test: %N, variance ratio ¼ 2.31, p ¼ 0.004; d15N, variance ratio ¼ 2.02, p ¼ 0.015) suggests that the d15N of sources for litter decay vary widely relative to those for wood decay fungi. The 15N enrichment of ectomycorrhizal fungi relative to saprotrophic fungi of about 5& is a well-established pattern across many studies (Mayor et al., 2009), but the underlying controls remain unclear. Here, saprotrophic fungi averaged 0.5
0.3&, hydrophilic ectomycorrhizal fungi averaged 3.2
0.2&, and hydrophobic ectomycorrhizal fungi averaged 8.6
0.4& (caps for all). The similar 15 N enrichments for caps versus stipes for saprotrophic fungi and for hydrophilic ectomycorrhizal fungi suggests similar internal processes in these two fungal groups that govern the partitioning of nitrogen isotopes in these fungi. Based on the 15N enrichments from the litter layer to deeper soil horizons that were measured here, the 15N enrichment of hydrophilic ectomycorrhizal taxa relative to saprotrophic taxa of w3& could arise solely from differences in source nitrogen. Observations in boreal forest by Lindahl et al. (2007) that saprotrophic fungi predominated in 15N-depleted surface litter whereas ectomycorrhizal fungi predominated at lower depths where d15N was higher also support interpreting 15N enrichments in hydrophilic ectomycorrhizal fungi relative to saprotrophic fungi as partially reflecting source differences. However, the transfer of 15 N-depleted N by ectomycorrhizal fungi to host plants has been

Fig. 4. Movement and isotopic fractionation of carbon isotopes in different components of plants, ectomycorrhizal fungi, and saprotrophic fungi. SAP ¼ saprotrophic, ECM ¼ ectomycorrhizal. CHO ¼ chitin and other carbohydrates. Protein (ho) indicates fungal protein from hydrophobic exploration types; protein (hi) indicates fungal protein from hydrophilic exploration types. Modified from Hobbie (2005).

shown in culture studies using isotopic mass balance to decrease plant d15N and increase fungal d15N (Hobbie and Colpaert, 2003) and must also contribute to the 15N enrichment of ectomycorrhizal fungi relative to saprotrophic fungi. 15 N enrichment of sporocarps, rhizomorphs, and mycelia of the hydrophobic ectomycorrhizal fungus Suillus relative to supplied N (Högberg et al., 1999b; Kohzu et al., 2000; Hobbie and Colpaert, 2003) and 15N enrichment of gills relative to stipes (Zeller et al., 2007) indicate that internal processes can create 15N differences in fungi even when the external source N is uniform. Accordingly, 15N partitioning among different compound classes must contribute to the high d15N of many hydrophobic taxa, with one mechanism the sequestration of relatively 15N-depleted nitrogen in chitin and wallbound protein in extraradical hyphae. Greater nitrogen sequestration in hydrophobic taxa than in hydrophilic taxa could lead to progressive 15N enrichment in the pool of internal nitrogen available for sporocarp formation. As depicted in Fig. 1b, 15N and 14N partitioning in a closed system could produce such enrichment patterns. 4.3. Explaining isotopic patterns in caps and stipes Differing compositions in caps and stipes of protein and carbohydrates should influence elemental and isotopic values in these fungal tissues. Taylor et al. (1997) attributed the higher %N and d15N in caps than in stipes to greater 15N-enriched protein and less 15 N-depleted chitin in caps than in stipes. In that study, caps were 2.4
0.3% higher in %N and about 2& enriched in 15N relative to stipes, and protein was 9.9
0.5& (n ¼ 6) higher in 15N than chitin, with a resulting calculated Dchitin of 9.8
0.5&. Here, we show that the 15N enrichments follow regular patterns and correlate with exploration type and the hydrophobicity of ectomycorrhizae. These are presumably important functional attributes that correlate with enzymatic activity, sensitivity to N deposition, and exploration depth (Hobbie and Agerer, 2010; Lilleskov et al., 2011; Peay et al., 2011). The few studies comparing the compositions of caps and stipes have focused on edible, saprotrophic species that are readily cultured. In Volvariella volvacea, protein content of caps was about twice that of stipes at the elongation stage, but differed less in later stages (Chang and Chan, 1973). Chitin content of caps was similar to that of stipes in Agaricus bisporus (chitincap/stipe ¼ 0.91) but significantly lower in two other species, Pleurotus ostreatus (chitincap/stipe ¼ 1.35) and Lentinula edodes (chitincap/stipe ¼ 1.20) (Vetter, 2007). In three Pleurotus species, protein was significantly higher (w60%) in caps than in stipes in three Pleurotus species but not in Calocybe indica, whereas carbohydrate content did not differ significantly between caps and stipes in any of these four species (Alam et al., 2008). In Lentinus subnudus, Psathyrella atroumbonata, and Termitomyces striatus, protein content doubled from stipes to caps whereas non-protein nitrogen content stayed roughly constant (cap concentration 90% that in stipes,

66

E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

calculated from Alofe et al., 1996). Based on these studies, we conclude that assuming a constant chitin content between caps and stipes is a reasonable simplification, with large increase in protein content from stipes to caps driving changes in %N and d15N. The 13C increase of caps relative to stipes presumably reflects higher 13C values of protein than in carbohydrates (Webb et al., 1998; A. Ouimette, pers. comm.). The 13C enrichment of protein relative to carbohydrates may primarily arise from the loss of 13C-depleted CO2 during respiration in the tricarboxylic acid (TCA) cycle (O’Leary and Yapp, 1978; Grissom and Cleland, 1988; Tcherkez et al., 2003), leaving a pool of 13C-enriched, TCA-derived carbon skeletons to construct many amino acids. The C:N ratio of protein is lower than the C:N of sporocarps (w3 versus w9; calculated from %C and %N data), therefore the difference in 13C between caps and stipes caused by differences in protein content is small relative to the 15N difference. In addition, protein is only several per mille higher in 13C than carbohydrates (Winkler et al., 1978; Webb et al., 1998) but up to 12& higher in 15N than chitin (Taylor et al., 1997; Webb et al., 1998). The higher calculated isotopic enrichment for both 15N and 13C in hydrophobic exploration types than in either hydrophilic exploration types or in saprotrophic fungi implies that patterns of internal sequestration and remobilization greatly influence 15N patterns and 13 C values in sporocarps. Studies that specifically measure isotopic patterns in fungal protein, carbohydrates (including chitin), and nucleic acids would help to further resolve these issues. 4.4. Explaining d15Ncapestipe through multiple regressions For two of the variables used in the multiple regression, 1  % Nstipe/%Ncap and %Ncap-stipe, multicollinearity is a potential problem. However, given the relatively modest level of multicollinearity (r2of the two variables is 0.50) and the very high sample size (n ¼ 286), it appears that the chance of this inducing substantial errors is very small (Mason and Perreault, 1991). We have accordingly retained both of these variables in the multiple regression, and interpret them separately. The multiple regression analysis presented in Table 2 confirms the importance of the relative nitrogen concentration in caps versus stipes (1  %Nstipe/%Ncap) in explaining d15Ncapestipe, as this value should correlate with the relative proportion of 15N-enriched protein and 15N-depleted chitin in these two fungal components. If shifts in % N between stipes and caps were only caused by adding protein to caps, with chitin content held constant, then the coefficient for this variable should equal the 15N enrichment of protein relative to chitin of 9.9
0.5& (Taylor et al., 1997), multiplied by the fraction of stipe nitrogen that is chitin (Equation (9)). Since 5.171&/9.9& ¼ 0.52, about half of the stipe nitrogen is chitin. Little data exist for comparison. Non-protein nitrogen was 33% of total nitrogen in whole mushrooms (Fujihara et al.,1995). Non-protein nitrogen in stipes was 51%, 17%, and 14% of total nitrogen for three saprotrophic species (calculated from Alofe et al., 1996), whereas 39% of stipe nitrogen was chitin in ectomycorrhizal fungi (calculations in Appendix 3 using data from Taylor et al., 1997). We expected that %Ncapestipe would positively correlate with d15Ncapestipe, given the higher %N in protein than in chitin, whereas the actual coefficient in the regression was negative (0.78&/%N). One plausible explanation is that stipes with low %N should average a greater proportion of nitrogen as chitin than stipes with high %N. If increases in the %N of caps relative to stipes are primarily attributed to added protein, then for a doubling of protein from stipes to caps, the relative shift in the fraction of nitrogen attributed to protein (and consequently, the shift in d15N) will be less for mushrooms with higher stipe %N. For example, with chitin at 6.89% N, protein at 16.7% nitrogen (calculated from the inverse of nitrogen:protein in Fujihara et al., 1995), and chitin at 8% of biomass, doubling protein from 7% to

14% from stipes to caps increases %N by 1.17% and increases d15N by 1.28&, whereas doubling protein from 14% to 28% increases %N by 2.33% and only increases d15N by 0.85&. For the values given here, the shift in d15Ncapestipe for increases in %Ncapestipe would be 0.63& per %N. The positive correlation of d13Ccapestipe with d15Ncapestipe reflects that the increased protein in caps relative to stipes is high in both 15 N and 13C. Given an assumed 15N enrichment of 9.9& (Taylor et al., 1997) and a calculated 13C enrichment of 4.2& (see first paragraph of next section), the expected coefficient is 2.36, if shifts in protein equally affect d15N and d13C. From the actual coefficient of 0.89, we calculate that the fraction of new carbon that is protein is 0.89/2.36, or 37.7%, of the fraction of new nitrogen that is protein. Given a C:N of protein of 2.86 (calculated from amino acid composition of four taxa in Mattila et al., 2002; calculations in Appendix 3), the estimated cap C:N is then 2.86/0.377, or 7.59, which is about 17% lower than the value of 9.17 calculated here from average %C (41.46%) and %N (4.49%). Having hydrophobic ectomycorrhizae increased d15Ncapestipe by 1.1& and accounted for about 15% of the explained variance, whereas having hydrophilic ectomycorrhizae decreased d15Ncapestipe by 0.6& and accounted for 7% of the explained variance. The other categorical variables (litter decay fungi or wood decay fungi) were not significantly correlated with d15Ncapestipe, suggesting that these fungi process nitrogen similarly between caps and stipes. The coefficient for these variables can be explained as reflecting variable 15N enrichment of protein relative to chitin among the four functional groups. If protein as a percentage of total nitrogen is 21% higher in caps than in stipes (calculated from Taylor et al., 1997), then the additional 15N enrichment in hydrophobic taxa of 1.084& in our regression reflects an additional enrichment of protein relative to chitin of 5.1&. Similarly, the coefficient in hydrophilic taxa of 0.571& reflects an additional enrichment of protein relative to chitin of 2.7&. If the baseline fractionation between protein and chitin is 9.9&, then the fractionation of chitin versus protein in hydrophobic taxa will be 15.0&, in hydrophilic taxa will be 7.2&, and in saprotrophic fungi will be 9.9&. The only comparable data are in Taylor et al. (1997), in which protein differed from chitin by 8.45& (Amanita, hydrophilic), 10.85&, and 10.3& (both Suillus species, hydrophobic). These differences in fractionation attributed to hydrophilic and hydrophobic ectomycorrhizal taxa suggest that patterns of nitrogen storage prior to sporocarp formation and nitrogen transfer to host plants can influence isotopic partitioning between caps and stipes, and presumably influence the underlying isotopic partitioning between fungal protein and chitin. 4.5. Explaining d13Ccapestipe through multiple regressions Correlations of d13Ccapestipe with %Ncapestipe could be attributed to the higher d13C and %N in protein than in chitin and other carbohydrates. If %N increases in caps because of added protein (assumed to be 16.67% nitrogen) and chitin content remains constant between caps and stipes, then the coefficient of %Ncapestipe, 0.253&/%N, can be used to estimate the 13C enrichment of protein relative to non-chitin components of 0.253
0.033& 16.67, or 4.2
0.5&. Carbohydrates dominate fungal composition, averaging 62% in four species (Mattila et al., 2002), and losses from stipes to caps to balance protein gains must necessarily be largely of carbohydrates (Alofe et al., 1996). Fungal carbohydrates and protein have yet to be compared isotopically; Webb et al. (1998) reported that locust protein was 1.6& enriched in 13C relative to chitin and 3.4& enriched in 13C relative to trehalose. The positive correlation of d13Ccapestipe with d15Ncapestipe reflects that the increased protein in caps relative to stipes is high in both 15 N and 13C. A coefficient of 0.42 is expected for the shift of

E.A. Hobbie et al. / Soil Biology & Biochemistry 48 (2012) 60e68

d13Ccapestipe with d15Ncapestipe if protein C/N and cap C/N are similar, since the increased protein in caps will be 4.2& higher in 13C and 9.9& higher in 15N than carbohydrates. The actual coefficient of 0.121
0.017 therefore suggests that cap C:N is 2.86  0.42/0.121, or 9.93, which is 8% higher than the actual value of 9.17, and 31% higher than the estimated C:N (7.59) from the coefficient of d13Ccapestipe when regressed against d15Ncapestipe. It is possible to combine information in the coefficient of d13Ccapestipe against d15Ncapestipe and the coefficient of d15Ncapestipe against d13Ccapestipe. We have expressed the coefficient of d13Ccapestipe as: 15 13 0:853 ¼ d Nproteinchitin =d Cproteinchitin  C=Nprotein =C=Ncap

(10) 15

13

For the coefficient of d Ncapestipe against d Ccapestipe, we have: 13 15 0:121 ¼ d Cproteinchitin =d Nptoteinchitin  C=Nprotein =C=Ncap

(11) Solving for C/Ncap gives:

C=Ncap ¼ C=Nprotein =ð0:853 x 0:121Þ0:5

(12)

For a C/Nprotein of 2.86, the estimated C/Ncap is then 8.90, which is within 3% of the value of 9.17 calculated from average %N and %C of caps. This close agreement suggests that our assumptions about the causes of shifts in d15N, d13C, and %N are internally consistent. The negative correlation between d13Ccapestipe and %Ccapestipe is presumably caused by variations in lipid content, as lipids are both higher in %C and lower in 13C relative to carbohydrates and protein (Poorter et al., 1997; Hobbie and Werner, 2004). Caps appear higher in lipid content than stipes in mushrooms (Alam et al., 2008), with three species in Alofe et al. (1996) averaging 3% lipids in stipes and 5% in caps. Information is lacking on the 13C depletion of fungal lipids relative to carbohydrates or protein in sporocarps, although phospholipid fatty acids were 2.4& and 7.4& depleted in 13C relative to bulk mycelia in two cultured taxa (calculated from Ruess et al., 2005). Although no categorical variables correlated with d13Ccapestipe, shifts in d13Ccapestipe caused by the categorical variables are effectively incorporated into the multiple regression via the coefficient of d15Nproteinechitin. That is, any shift in d15Nproteinechitin caused by being hydrophobic or hydrophilic will then change d13Ccapestipe according to the given coefficient (0.121). If an additional factor was altering d15N that was not altering d13C, then the estimated C/Ncap would have not agreed so closely with the measured value. We conclude that processes differing in magnitude between hydrophobic and hydrophilic taxa that alter nitrogen isotopes also alter carbon isotopes. 4.6. Conclusions Patterns of nitrogen mobilization and isotopic fractionation between protein and other nitrogen-containing molecules appear to be the main driver of the 15N enrichment of caps relative to stipes. Although the fractionation is framed as reflecting two main pools in fungi, protein and chitin, other nitrogen-containing molecules could also contribute, such as nucleic acids (Gottlieb and Van Etten, 1964; Iten and Matile, 1970). For example, Fujihara et al. (1995) estimated that the relative contributions of different nitrogen forms to sporocarps in 13 species varied from 67% (protein) to 2% (NH3), with intermediate amounts for chitin (9%), nucleic acids (6%), and unknown nitrogen (16%). Chikaraishi et al. (2007) suggested that 15N depletion during aminotransferase reactions (such as movement of an amido group from glutamine to glucose to form glucosamine)

67

drive many d15N patterns during amino acid metabolism. It is possible that such aminotransferase reactions control the 15N depletion of both fungal chitin and ectomycorrhizal plants relative to fungal protein. If nitrogen were delivered as a single isotopically uniform source to sporocarps and then differentiated into protein or chitin, we would expect that 15N differences between caps and stipes would be similar across exploration types when corrected for differences in %N. Instead, our results suggest two non-exclusive possibilities: (1) the two main nitrogen pools in sporocarps, chitin and protein, can have different internal sources and are delivered separately to the developing primordia, and (2) source nitrogen is processed differently in hydrophobic ectomycorrhizal fungi relative to other fungal types into protein and chitin. The mechanistic and conceptual explanations offered here on the causes of isotopic differences in sporocarps are unlikely to be the full story. However, we hope that these ideas will stimulate further critical research and analyses of the underlying mechanisms causing these patterns, and therefore provide additional insights into the belowground functioning in carbon and nitrogen dynamics of ectomycorrhizal and saprotrophic fungi. One promising avenue is indicated by the existing literature on fungal metabolism during sporocarp formation (Moore, 1998). Such studies have revealed extensive differences in enzymatic processes between caps and stipes; for example, activities of NADP-glutamate dehydrogenase, glutamine synthetase, ornithine acetyltransferase, and ornithine carbamyltransferase are higher in the cap than the stipe of Coprinus cinereus fruitbodies while urease activity is much higher in stipes (Moore, 1998). The reactions catalyzed by these enzymes will influence the movement of amino acids, ammonia, and other compounds that ultimately control nitrogen and carbon isotope distributions. Studies under controlled conditions in which isotopic patterns of stipes, caps, protein, and chitin could be compared against differences in composition and metabolic pathways would also be informative. Acknowledgements We thank John Hobbie, Luke Nave, Andy Ouimette, Don Phillips, Stephen Trudell, and two anonymous reviewers for comments on prior versions. The US Environmental Protection Agency partially funded this research, which has been reviewed and approved for publication as an EPA document. Mention of trade names or commercial products does not constitute endorsement or recommendation for use. This work was also supported by two grants from the U.S. NSF Division of Environmental Biology, by a research fellowship to F.S. by the Scientist Committee of NATO, and by a Bullard Fellowship to E.H. from Harvard University. Appendix. Supplementary material Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.soilbio.2012.01.014. References Agerer, R., 2001. Exploration types of ectomycorrhizae e A proposal to classify ectomycorrhizal mycelial systems according to their patterns of differentiation and putative ecological importance. Mycorrhiza 11, 107e114. Agerer, R., 2006. Fungal relationships and structural identity of their ectomycorrhizae. Mycological Progress 5, 67e107. Alam, N., Amin, R., Khan, A., Ara, I., Shim, M.J., Lee, M.W., Lee, T.S., 2008. Nutritional analysis of cultivated mushrooms in Bangladesh e Pleurotus ostreatus, Pleurotus sajor-caju, Pleurotus florida and Calocybe indica. Mycobiology 36, 228e232. Alofe, F.V., Odeyemi, O., Oke, O.L., 1996. Three edible wild mushrooms from Nigeria: their proximate and mineral composition. Plant Foods for Human Nutrition 49, 63e73.

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