Research
Dead fungal mycelium in forest soil represents a decomposition
hotspot and a habitat for a specific microbial community
Vendula Brabcova, Monika Nova kova , Anna Davidova and Petr Baldrian
Institute of Microbiology of the ASCR, v.v.i., Vıde
nska 1083, 14220, Praha 4, Czech Republic
Summary
Author for correspondence:
Vendula Brabcov
a
Tel: +420 774 358 455
Email: [email protected]
Received: 12 November 2015
Accepted: 7 December 2015
New Phytologist (2016) 210: 1369–1381
doi: 10.1111/nph.13849
Key words: bacteria, decomposition,
enzyme activity, fungi, mycelium turnover,
soil, temperate forest.
Turnover of fungal biomass in forest litter and soil represents an important process in the
environment. To date, knowledge of mycelial decomposition has been derived primarily from
short-term studies, and the guild of mycelium decomposers has been poorly defined.
Here, we followed the fate of the fruiting bodies of an ectomycorrhizal fungus in litter and
soil of a temperate forest over 21 wk. The community of associated microbes and enzymatic
processes in this specific substrate were described.
The decomposition of fungal fruiting bodies exhibited biphasic kinetics. The rapid initial
phase, which included the disappearance of DNA, was followed by a slower turnover of the
recalcitrant fraction. Compared with the surrounding litter and soil, the mycelium represented
a hotspot of activity of several biopolymer-degrading enzymes and high bacterial biomass.
Specific communities of bacteria and fungi were associated with decomposing mycelium.
These communities differed between the initial and late phases of decomposition. The bacterial community associated with decomposing mycelia typically contained the genera
Pedobacter, Pseudomonas, Variovorax, Chitinophaga, Ewingella and Stenotrophomonas,
whereas the fungi were mostly nonbasidiomycetous r-strategists of the genera Aspergillus,
Penicillium, Mortierella, Cladosporium and several others.
Decomposing ectomycorrhizal fungal mycelium exhibits high rates of decomposition and
represents a specific habitat supporting a specific microbial community.
Introduction
In a forest ecosystem, fungi play key roles in nutrient cycling as
a consequence of both the significant involvement of saprotrophic taxa in the degradation of dead plant biomass (de Boer
et al., 2005; Osono, 2007; Baldrian & Valaskova, 2008) and
the mediation of carbon (C) flow from plant roots into the soil
by mycorrhizal taxa (Clemmensen et al., 2013; van der Heijden
et al., 2015). Fungal mycelia represent an important pool of
organic matter in forest litter and soil (Baldrian et al., 2013b;
Soudzilovskaia et al., 2015). Estimates of mycelial biomass for
ectomycorrhizal (ECM) fungi, which represent the bulk of soil
fungal biomass, typically range from 100 to 600 kg ha1 (Wallander et al., 2004; Cairney, 2012; Hendricks et al., 2016).
More importantly, yearly mycelial production is estimated to
occur at the same order of magnitude. For example, in a review
by Ekblad et al. (2013), the annual production of fungal
mycelia in Picea abies forests in the upper 10 cm of soil typically
ranged between 100 and 300 kg ha1 yr1. Significant amounts
of mycelia are also produced in soil layers below a depth of
10 cm (Wallander et al., 2004; Bostrom et al., 2007; Majdi
et al., 2008) and in litter, where the content of fungal biomass
per g substrate dry mass can be > 10-fold that of soil (Baldrian
et al., 2013b).
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Although a small fraction of the nutrients that constitute fungal mycelia may be retained in the soil and contribute to C storage by forest ecosystems, the decomposition of mycelia clearly
represents an important process for nutrient cycling in forest
soils. The dead fungal mycelia typically contain mostly cell wall
materials, such as polysaccharides, which represent 80–90% of
the total cell wall mass, lipids and proteins (Baldrian et al.,
2013b). Three to sixty per cent of the polysaccharide fraction is
composed of chitin and various other types of glucans, glucomannans and other polysaccharides containing galactose, galactosamine,
fucose or other components (Bartnicki-Garcia, 1968; Nilsson &
Bjurman, 1998). The amount of chitin and protein in fungal cell
walls is relatively high, and their biopolymers thus represent an
important source of both C and nitrogen (N) (Cooke & Whipps,
1993; Colpaert et al., 1996; Wallander et al., 2004; Zeglin &
Myrold, 2013). The C : N ratio of dead fungal biomass can be as
low as 7, whereas the average ratio of fresh leaf litter is 25–200
(Valaskova et al., 2007; Koide & Malcolm, 2009; Mouginot et al.,
2014). The high content of N thus makes fungal mycelia attractive
targets of decomposition in forest litter and soil, which are often N
limited (Lindahl et al., 2007; Snajdr et al., 2008).
The attractiveness of fungal mycelia as targets of decomposition is demonstrated by their initial rates of decomposition,
which are, despite variation among fungal taxa, typically faster
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than the initial rates of litter decomposition. For example, in a
wide range of tree litters, 7–50% of the mass was lost within the
first 6 months (Osono & Takeda, 2005), whereas mass loss of an
equal extent only required 4 wk in ECM fungal mycelia (Koide
& Malcolm, 2009). Mycelia with higher chitin or N content
decompose more quickly (Fernandez & Koide, 2012, 2014) and
their decomposition is not retarded by presence of recalcitrant
lignin, which typically slows the decomposition of plant litter
(Berg, 2000; Snajdr et al., 2011), although the presence of melanin
in the walls of certain fungi may have the same effect (Fernandez
& Koide, 2014). Unfortunately, the existing studies on mycelium
decomposition are limited to short periods of time and do not consider the longer term fate of the mycelial biomass. Recently, an
important finding suggested that the majority of C stored in deeper
horizons of boreal forest soils originates from the allocation of photosynthate to fungi in symbiosis with tree roots (Clemmensen
et al., 2013) and that certain mycelial fractions, such as those rich
in melanin, can be recalcitrant to decomposition (Koide & Malcolm, 2009; Ekblad et al., 2013), which indicates that the rate of
decomposition may be lower after an initial rapid phase.
Considering the quantitative importance of dead fungal
mycelia, surprisingly little is known about the microorganisms
that decompose it. Although the functional guilds of fungi that
form symbiotic mycorrhizal associations with plant roots or act as
litter decomposers are well established (Tedersoo et al., 2014;
Clemmensen et al., 2015) and there has been some progress in
understanding bacterial involvement in plant litter decomposition
(Stursova et al., 2012; Berlemont & Martiny, 2015), the guild of
mycelium decomposers, if it exists, has not yet been described in
the environmental context. The mycophagous activity of selected
model species of fungi and bacteria, for example Trichoderma,
Collimonas, Paenibacillus, Pseudomonas and Myxobacterium, has
been extensively described (de Boer et al., 2005; Guthrie &
Castle, 2006; Leveau & Preston, 2008; Seidl, 2008; Cao et al.,
2009; Ihrmark et al., 2010), but all of the studies performed to
date in an environmental context have been indirect. Measurements obtained by inserting soil sampling tubes indicate that certain free-living ascomycetes increased in soil cores shortly after the
roots and mycelia were severed, although the bacterial populations seemed little affected. These results indicate that such disturbances may induce the rapid growth of opportunistic
saprotrophic fungi that presumably use dead mycorrhizal
mycelium along with other resources (Lindahl et al., 2010). In
another study, free-living soil fungi accumulated C from labelled
ECM fungal mycelia, which suggested that basidiomycete fungi
play a role in the process (Drigo et al., 2012) but left the respective roles of different fungi undetermined.
The aim of this study was to follow in detail the decomposition of dead fungal mycelium originating from whole fruiting
bodies of an ECM fungus in a temperate deciduous forest over
an extended period of time and to link data on the kinetics of
decomposition, chemical composition and activities of extracellular enzymes with a description of the associated microbial community. To accomplish this, bags with fungal mycelia were
incubated in situ and analysed periodically. Because the communities of microorganisms that inhabit the litter and soil differ
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(Vorıskova et al., 2014; Lopez-Mondejar et al., 2015), the study
was performed in both of these compartments. Based on the
chemical composition of mycelia, we hypothesized that dead fungal mycelium, at least in the initial stages of decomposition,
could support high microbial activity, supporting more biomass
than the rest of the litter or soil material. However, the initial
rates of mycelium turnover are unlikely to remain high over prolonged incubation, and we hypothesized that a substrate-driven
succession of functionally different decomposers would occur, as
occurs in plant litter (Vorıskova & Baldrian, 2013).
Materials and Methods
Study site, materials and experimental set-up
The study site was a temperate oak (Quercus petraea (Matt.)
Liebl) forest in the Xaverovsky Haj Natural Reserve, near Prague,
Czech Republic (50°150 38″N, 14°360 48″E). The soil was an
acidic cambisol with a developed litter and organic and mineral
horizons (Baldrian et al., 2013a). The site has been previously
studied with respect to the activity of decomposition-related
extracellular enzymes (Snajdr et al., 2008; Baldrian et al., 2010,
2013a; Vetrovsky et al., 2013) and the decomposition of leaf litter and associated changes in the community of fungal decomposers (Snajdr et al., 2011; Vorıskova & Baldrian, 2013).
Importantly, the microbial community composition in the forest
topsoil and its changes across seasons have also been described
(Vorıskova et al., 2014; Lopez-Mondejar et al., 2015). The content of fungal biomass at the study site ranges typically between
10 and 20 mg g1 in litter and 0.8–2.6 mg g1 in the soil organic
horizon (Snajdr et al., 2008; Baldrian et al., 2013b).
Fungal biomass was obtained by collecting the fruiting bodies
of the ECM fungus Tylopilus felleus (Bull.) P.Karst., which is a
species associated with Picea abies (L.) H.Karst. The species does
not occur at or near the study site, so its presence in samples did
not interfere with the DNA-based analyses. Whole fresh fruiting
bodies were cut into 4-mm pieces, freeze-dried and stored at
room temperature. Mycobags – mesh bags (10 9 20 cm; 1 mm
polyester mesh size) that were filled with 3 g of mycelia – were
sterilized by repeated gamma-irradiation and placed in the
middle of each layer (thickness 1–2 cm) and the soil organic horizons (thickness 1.5–3 cm). The experiment started in the early
spring, and the mycobags were removed after 1, 2, 3, 9, 15 and
21 wk of incubation. Four mycobags incubated in litter and four
incubated in soil were collected at each time-point, along with
four litter and soil samples from the same depth, but different
randomly selected locations within 1 m of the mycobags, which
were used as background controls. Material was transferred to the
laboratory, cut if necessary, freeze-dried and stored at 20°C.
Sample chemistry and enzyme activity
Loss of dry mass was measured after freeze-drying, the organic matter content was estimated after combustion at 650°C, and soil pH
was measured in distilled water (1 : 10 w/vol) after 2 h. Oxidizable
C (Cox) and total N contents were measured using an elemental
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analyser (Elementar Vario EL III, Elementar, Hanau, Germany) in
an external laboratory (Central laboratory of University of the
Chemistry and Technology, Prague, Czech Republic). Cox was
measured using sulfochromic oxidation, and N content was estimated by the Kjeldahl method (Bremner, 1960).
The activities of extracellular enzymes were assayed in sample
homogenates as described previously (Stursova & Baldrian,
2011). Briefly, the activities of b-glucosidase, a-glucosidase, cellobiohydrolase
(exocellulase),
b-xylosidase,
Nacetylglucosaminidase, b-mannosidase, a-galactosidase, lipase,
phosphomonoesterase (phosphatase) and alanine aminopeptidase
were measured at pH 5.0 in 1 : 50 (w/v) soil slurries using methylumbelliferol and amidomethylcoumarin-based substrates on a
microplate reader (Infinite 200 PRO, Tecan, Mannedorf,
Switzerland), with an excitation wavelength of 355 nm and an
emission wavelength of 460 nm. Calibration of product development was based on standard curves with a range of 4-methylumbelliferyl (MUF) and 7-amino-4-methyl coumarin (AMC)
concentrations in the same soil slurry.
DNA extraction, microbial biomass quantification and
amplicon sequencing
Total DNA was extracted from 300 mg of sample material using
a modification of the method of Miller based on phenolchloroform extraction (Sagova-Mareckova et al., 2008) and
cleaned with a GeneClean Turbo Kit (MP Biomedicals, Solon,
OH, USA). Bacterial and fungal rRNA gene copies were quantified by qPCR using the 1108f and 1132r primers for bacteria
(Wilmotte et al., 1993; Amann et al., 1995) and FR1 and FF390
primers for fungi (Vainio & Hantula, 2000; Prevost-Boure et al.,
2011). Plasmids containing amplified fragments of Streptomyces
lincolnensis DNS 40335 and Hypholoma fasciculare CCBAS 286
were used as standards. Three technical replicates were performed
for each sample, and all samples from all four replicates for each
treatment were sequenced.
The general bacterial primers eub530F/eub1100aR (Dowd
et al., 2008) were used to amplify the V4–V6 region of bacterial
16S rDNA and the fungus-specific primers ITS1/ITS4 (White
et al., 1990) were used to amplify the ITS1, 5.8S rDNA and
ITS2 regions of fungal rDNA as described previously (Baldrian
et al., 2012). Briefly, two-step PCR amplification using barcoded
primers was performed with three independent PCR reactions
per sample. PCR products were purified using Agencourt
AMPure XP (Beckman Coulter, Beverly, MA, USA) and an
equimolar pool of PCR products from all samples was prepared.
After gel electrophoresis, the pool was purified using the Wizard
SV Gel and PCR Clean-Up System (Promega, Madison, WI,
USA), the Agencourt AMPure XP and a MinElute PCR Purification Kit and sequenced by 454-pyrosequencing.
Bioinformatics analysis and statistics
The pyrosequencing data were processed using the pipeline SEED
1.1.2 (Vetrovsky & Baldrian, 2013). Pyrosequencing noise
reduction was performed using DENOISER 0.851 (Reeder &
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Knight, 2010), and chimeric sequences were detected using
UCLUST 3.0 (Edgar et al., 2011) and deleted. Sequences shorter
than 380 bases were removed. All remaining sequences were
shortened to 380 bases and clustered using USEARCH 5.2 (Edgar,
2010) at a 98% similarity level. Consensus sequences were constructed for each cluster, and the operational taxonomic units
(OTUs) were constructed by clustering these consensus
sequences at 97% identity (Lundberg et al., 2012). The abundance data reported in this paper are based on this data set of
sequence abundances and should be taken as proxies of taxon
abundance with caution (Lindahl et al., 2013). The closest hits
to fungal and bacterial consensus sequences were identified by
comparison with the UNITE database (Koljalg et al., 2013) and
the Ribosomal Database Project (Cole et al., 2009), respectively.
Fungal genera were assigned a predicted ecophysiology based on
Tedersoo et al. (2014). Sequence data were deposited in MG
RAST (Meyer et al., 2008) (data set numbers 4676034.3 and
4676035.3).
Diversity estimates (Shannon–Wiener index, species richness
and evenness) were calculated for a data set containing 1000 randomly chosen sequences from each sample in SEED 1.1.2. (Vetrovsky & Baldrian, 2013). Statistical analyses were performed using
the software package STATISTICA 7 (StatSoft, Tulsa, OK, USA). In
the nonmetric multidimensional scaling analysis, only OTUs
with relative abundances ≥ 1.0% in at least three samples were
included (82.3% and 81.75% of all bacterial and fungal
sequences, respectively) to reduce the extent of random variation.
The significance of differences in community composition was
determined using analysis of similarities (ANOSIM) with Bray–
Curtis similarity, and nonmetric multidimensional scaling was
used to visualize the treatment effects. The significance of differences in the relative abundances of individual bacterial and fungal
OTUs and genera, enzyme activities and nutrient contents were
determined using ANOVA and the Fisher least significant
difference post hoc test. Differences with P < 0.05 were considered statistically significant.
Results
Decomposition of fungal mycelia
The mass loss of fungal mycelia during the first 2 wk was very
rapid, reaching 0.21 wk1. It later slowed and was very slow at
the end of the experiment (0–0.06 wk1). The decomposition of
the mycelium was significantly faster in soil than in litter at the
beginning of the experiment, but the remaining mass of mycelia
after 21 wk was equal at 25% (Fig. 1). Transformation of the
substrate was also apparent; the relative abundance of DNA reads
of Tylopilus felleus decreased in the mycobags and they virtually
disappeared within 9 wk when mycobags were incubated in litter
and within 21 wk when they were incubated in soil (Fig. 1).
The organic matter content was 94.4 0.0% in mycelia,
91.5 0.2% in litter and 30.2 2.2% in soil at the beginning of
the experiment, and the initial contents of N , C and hydrogen
were significantly higher in mycelia (5.1% N; 44.2% C) than in
litter (2.0% N; 40.1% C) and soil (1.5% N; 28.4% C). The
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(a)
(b)
90
120
Loss of mycelium dry mass
Tylopilus felleus DNA sequence abundance
Loss of dry mass (%)
70
60
50
Mycobags in soil
40
Mycobags in litter
30
20
Relative abundance (%)
80
Mycobags in soil
Soil
Mycobags in litter
er
100
80
60
40
20
10
0
0
0
5
10
15
20
25
0
5
10
Time (wk)
(c)
20
25
(d)
200
250
Mycobags in soil
Soil
Mycobags in litter
er
160
140
120
100
80
60
40
Copies g–1 dry mass (×10 7)
16S rDNA gene copies
180
Copies g–1 dry mass (×10 7)
15
Time (wk)
18S rDNA gene copies
Mycobags in soil
Soil
Mycobags in litter
er
200
150
100
50
20
0
0
0
5
10
15
20
25
Time (wk)
0
5
10
15
20
25
Time (wk)
Fig. 1 (a) Loss of mycelial dry mass, (b) relative abundance of Tylopilus felleus reads, (c) bacterial abundance and (d) fungal abundance during the
decomposition of fungal mycelia in forest litter and soil. The data represent means of four replicates with SE. Values for the control litter and the soil are
expressed as the means of estimates at all sampling times.
initial C : N ratio in mycelia (8.7) was significantly lower than
those in the surrounding litter (19.9) and soil (19.1). The C : N
ratio in the mycobags decreased slightly but significantly during
decomposition to 7.4 within 21 wk (P < 0.003). The initial pH
of the mycelia was 5.5, compared with 5.1 in litter and 4.3 in
soil. During the experiment, the pH in the mycobags increased
to 6.6 in litter, but it decreased to 5.3 in soil.
High enzyme activities in the mycobags had already been
recorded before the experiment (week 0), which probably represented the enzymes of T. felleus. The initial phase of decomposition was characterized by rapid changes in enzyme activities in
the mycobags so that the activity after 3 wk of decomposition was
significantly different from the initial activity for most enzymes
(Fig. 2). By contrast, the activity of enzymes showed relatively little change during the late phase of decomposition during weeks
9–21 (Fig. 2). Compared with week 0, the activities of
b-glucosidase, b-xylosidase, b-mannosidase, a-galactosidase and
alanine aminopeptidase decreased, whereas those of lipase and
phosphatase increased at least temporarily and the activity of Nacetylglucosaminidase remained relatively stable (Fig. 2).
Throughout the experiment, the activity of most enzymes in the
mycobags was significantly higher than the activity observed in
the litter or soil environment. Compared with litter, the enzyme
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activity in the mycobags incubated in litter during the late phase
of decomposition (weeks 9–21) was most increased for
N-acetylglucosaminidase (179), a-glucosidase (49) and
b-mannosidase (49). The differences were even greater when
comparing the mycobags incubated in soil with the soil: the difference was 26-fold for b-mannosidase, 21-fold for a-glucosidase,
21-fold for cellobiohydrolase, 20-fold for N-acetylglucosaminidase,
and 11-fold for both b-glucosidase and phosphatase (Fig. 2).
Bacteria associated with decomposing fungal mycelia
Bacterial abundance expressed as copies of 16S rDNA gradually
increased and reached the same abundance as in the surrounding
litter or soil only after 3 wk of incubation. In the remaining time,
bacterial abundance in the mycobags was significantly higher
than that in the controls, with a peak at week 15 in litter
(1.4 9 109 copies g1; four-fold higher than the litter average)
and at week 9 in soil (1.0 9 109 copies g1; 12-fold higher than
in the surrounding soil; Fig. 1).
Overall, 138 944 sequences of the bacterial 16S rDNA were
retained for analysis after denoising and removing both chimeric
and nonbacterial sequences (< 0.5%). Bacterial sequences clustered into 5563 OTUs at a 97% similarity level. The OTUs with
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400
60
β-glucosidase
α-glucosidase
a
Soil
350
a
a
Lier
Soil
50
Lier
300
40
250
a
b
200
a
30
b
150
c
20
100
b
c
10
50
c
b
0
0
Week 3
0
120
a
a ab
b
Weeks 9-21
Control
Cellobiohydrolase
Soil
100
a
Week 3
Weeks 9-21
Control
β-xylosidase
Soil
60
Lier
a
a
0
70
Lier
a
a
50
80
40
60
b
30
40
b
b
b
ab
b
b
20
c
c
20
0
0
0
200
c
10
b
Week 3
Weeks 9-21
Control
N-acetylglucoseaminidase
a
a
a
Soil
ab
Lier
150
0
400
b
Week 3
Weeks 9-21
Control
β-mannosidase
350
Soil
a
a
Lier
300
b
250
b
200
100
b
150
100
50
c
c
0
1200
c
50
c
c
c
0
0
Week 3
Weeks 9-21
Control
140
Phosphatase
Soil
a
1000
0
a
120
Week 3
Weeks 9-21
Control
α-galactosidase
a
Soil
a
Lier
Lier
100
800
a
ab
b
b
ab
80
600
60
b
b
400
40
200
b
b
20
b
c
c
0
0
0
900
Fig. 2 Activities of extracellular enzymes
during the decomposition of fungal mycelia
of Tylopilus felleus in forest litter and soil.
The data represent the means SE estimated
in the mycobags at weeks 0 and 3, in all
mycobags collected in weeks 9–21 and in
litter and soil sampled at all sampling times
(controls). Statistically significant differences
are indicated by different letters.
Week 3
Lipase
a
0
Control
Week 3
a
Soil
45
700
Lier
40
Control
Soil
a
Lier
a
35
600
b
b
500
30
25
bc
400
20
300
200
Weeks 9-21
50
800
c
c
c
b
15
b
10
100
5
0
0
0
relative abundance ≥ 1% in at least three samples are listed in
Supporting Information Table S1.
The bacterial community in the mycobags differed significantly from the background community in soil and litter at
all sampling times (ANOSIM on Bray–Curtis distances with
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Weeks 9-21
Week 3
Weeks 9-21
Control
b
bc
b
0
Week 3
Weeks 9-21
c
Control
9999 permutations: P = 0.007 in soil and P = 0.005 in litter).
Communities of bacteria at different sampling times were significantly different during the early (1–3 wk) and late (9–
21 wk) phases of decomposition. The differences in the bacterial community composition among mycobags of the same
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age from different horizons were small, and the decomposition
time was the most important factor governing the bacterial
community, whereas the location of the mycobags had a
smaller effect (Fig. 3).
The bacterial community in the litter in which the mycobags
were placed consisted primarily of Proteobacteria, Bacteroidetes
and Acidobacteria; the soil was dominated by Acidobacteria, Proteobacteria and Actinobacteria (Fig. 4). In the mycobags until
week 3, the bacterial communities were largely dominated by
Proteobacteria, especially Gammaproteobacteria, and the late
phases of decomposition were dominated by the Bacteroidetes
(up to > 80%). Actinobacteria increased towards the end of the
experiment, but the abundance of their sequences only reached
4% (Fig. 4). The genera Telmatobacter, Acidopila, Granulicella
and Candidatus Koribacter were the most common in litter, and
Mucilaginibacter, Burkholderia, Luteibacter and Granulicella were
the most common in soil (Fig. 5; Table S2). By contrast, decomposed mycelia were highly dominated by only a few taxa, most
notably Pedobacter (39% and 33% on average in the mycobags in
soil and litter, respectively) and Pseudomonas (34% and 23% in
soil and litter, respectively). Additionally, the bacterial community in the mycobags incubated in soil was significantly enriched
by Ewingella, Stenotrophomonas, Mucilaginibacter, Erwinia and
Ochrobactrum, whereas the mycobags incubated in litter showed
enrichment by Chitinophaga and Variovorax (Fig. 5).
The successive development of the bacterial community in
mycobags could be separated into an early and a late phase:
Pseudomonas and Ewingella dominated in the early phase of
decomposition but remained abundant later, whereas the abundance of Pedobacter increased with time. The abundances of
0.15
(a)
additional genera increased towards the end of the experiment,
specifically Variovorax and Chitinophaga in mycobags incubated
in the litter and Stenotrophomonas and Sphingobacterium in
mycobags incubated in the soil (Fig. 5). Compared with the surrounding soil and litter, the bacterial communities in mycobags
were significantly less diverse (P < 0.0001). The OTU richness at
1000 sequences/sample was on average 412 17 (mean SE) in
soil and 396 25 in litter, but only ranged from 50 to 200 in the
mycobags. In the mycobags, the relatively high species richness at
week 1 (150–200) dropped to < 50 at week 3 and then gradually
increased until it reached 130–150 at week 21. The mycobagassociated communities also showed much lower evenness (0.5–
0.7 compared with 0.9 in litter and soil, respectively; P < 0.0001).
Fungi associated with decomposing fungal mycelia
The abundance of fungal DNA in the mycobags was affected
because they contained T. felleus DNA (Fig. 1). The counts of
18S rDNA genes decreased during the experiment to a minimum
during weeks 9–15 (1.0–1.6 9 108 copies g1). Compared with
the surrounding substrate, the mycobags contained more 18S
rDNA copies when incubated in soil but substantially fewer
copies when incubated in litter after week 9 (Fig. 1).
Overall, 129 046 sequences of fungal internal transcribed
spacer (ITS) were used for analysis after removing the sequences
that were chimeric or nonfungal, or belonged to T. felleus
(27.5%). Because of the dominance of T. felleus in the DNA pool
at week 1, fungal community analysis in the mycobags incubated
in litter was only possible for weeks 2–21. After removal of
T. felleus sequences, fungal sequences clustered into 2365 OTUs
0.25
(b)
Fungi
Bacteria
2
0.2
0.1
0
21
0.05
21
0
0
21
21
21
0
0.15
2
2
0.1
3
0
3
3
3
0
1
–0.05
1
1
2
3
2
99
9
21
–0.05
2
er
2
1
2
2
3
2
–0.15
–0.1
15
–0.05
15
21
21
0
21
15
0
21
21
15
21
21
0
0.05
21
21
0
–0.15
Soil
0
0
Mycelium soil
3
0.1
21
Mycelium litter
2
0
0
21
er
2
1
Mycelium soil
15
21
–0.1
1
Mycelium litter
–0.2
2
21
21
21
9 15
0
3
–0.15
–0.25
3
15
1
2
Soil
9
3
3
–0.1
9
15
L0
21
9 9
9
0
2
1
3
2
1
3
1
3
0.05
21
0
0
21
9
9
21 15
21 15
21 9
21
15
21
15
9
21
15
21
15 15
21
9
15
9
9
0.15
–0.2
–0.25 –0.2 –0.15 –0.1 –0.05
0
0.05
0.1
0.15
0.2
0.25
0.3
Fig. 3 Nonmetric multidimensional scaling of (a) bacterial and (b) fungal community composition during the decomposition of fungal mycelia of Tylopilus
felleus in forest litter and soil, based on operational taxonomic units with relative abundance ≥ 1% in at least three samples. Numbers indicate weeks of
sampling.
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100%
(a)
90%
Relative abundance
80%
70%
60%
50%
Gammaproteobacteria
Betaproteobacteria
Alphaproteobacteria
Deltaproteobacteria
Bacteroidetes
Acnobacteria
Firmicutes
Acidobacteria
Verrucomicrobia
Gemmamonadetes
Cyanobacteria
Chlamydiae
Armamonadetes
Chloroflexi
Fusobacteria
Planctomycetes
Tenericutes
Other
40%
30%
20%
10%
0%
100%
(b)
90%
Relative abundance
80%
70%
60%
50%
40%
30%
20%
10%
0%
Control
Week 1
Week 2
Week 3
Week 9
Week 15
Week 21
100%
(c)
90%
Relative abundance
80%
70%
Euroales
Capnodiales
Leoomycetes_other
Heloales
Hypocreales
Ascomycota_other
Glomerellales
Lecanoromycetes_other
Dothideomycetes_other
Xylariales
Saccharomycetales
Pleosporales
Morerellales
Basidiomycota_other
Tremellales
Thelephorales
Boletales
Russulales
Agaricales
Polyporales
Mucorales
Rhizophydiales
Other Fungi
60%
50%
40%
30%
20%
10%
0%
100%
(d)
90%
Relative abundance
80%
70%
60%
50%
40%
30%
20%
10%
0%
Control
Week 1
Week 2
Week 3
Week 9
Week 15
Week 21
Fig. 4 Composition of the bacterial community associated with decomposing fungal mycelia of Tylopilus felleus in (a) forest litter and (b) forest soil, sorted
into phylum level (and into class level for proteobacteria) and composition of fungal community associated with decomposing fungal mycelia in (c) forest
litter and (d) forest soil, sorted into order level. The data represent means (n = 4) of relative abundances.
at the 97% similarity level. The OTUs with a relative abundance
≥ 1% in at least three samples are listed in Table S3.
The fungal community in the mycobags differed significantly
from those in the surrounding soil and litter (ANOSIM on
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Bray–Curtis distances with 9999 permutations: P = 0.006 in soil
and P = 0.005 in litter). Importantly, the communities differed
significantly between mycobags incubated in the litter and in the
soil at all sampling times (P = 0.032). Although the early and late
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Week 15
Week 21
Week 9
Week 15
Week 21
10
100
Week 3
Week 9
Week 3
1.0
Week 2
Week 2
0.1
Soil
0
Week 1
Soil
Week 1
Week 21
Week 15
Week 3
Week 9
Week 1
Litter
Syntrophaceticus
Candidatus Solibacter
Rhodopila Actinoallomurus Edaphobacter
Acidobacterium Acidopila
Granulicella
Frankia Acidisphaera Aciditerrimonas
Actinomadura Acidocella Telmatobacter
Burkholderia Iamia Phormidium
Bryobacter
Methylosinus Ideonella Candidatus Koribacter
Ferruginibacter Ewingella Conexibacter
Luteibacter Pedosphaera
Mucilaginibacter Pseudomonas Azospirillum Lysobacter
Frondihabitans
Terriglobus Bradyrhizobium Novosphingobium Mycobacterium Erwinia
Sphingomonas Janthinobacterium Chthoniobacter +
Sphingobacterium +
Phenylobacterium +
Paenibacillus +
Achromobacter +
Pedobacter +
Variovorax +
Rhizobium +
Pseudochrobactrum +
Stenotrophomonas +
Ochrobactrum +
Devosia +
Chitinophaga +
Rhodococcus +
Soil
Sympodiella
Odonticium
Troposporella
Mycena
Cylindrosympodium
Phyllosticta
Penicillium
Gloiodon
Athelia
Dwayaangam
Mortierella
Devriesia
Cryptococcus
Lachnellula
Cadophora
Cryptosporiopsis
Aspergillus
Meliniomyces
Cladosporium
Phialocephala
Mycolevis
Fulvoflamma
Pseudeurotium
Trichosporon
Lactarius
Geomyces
Russula
Crocicreas
Oidiodendron
Leptodontidium
Xerocomus
Tomentella
Epicoccum
Mucor
Rinodina
Pseudaegerita
Cenococcum
Elaphomyces
Campylocarpon
Pochonia
Mycoleptodiscus
Candida
Asterotremella
Fusarium
Calcarisporium
Kappamyces
Monographella
Verticillium
Edaphobacter Frankia Acidisphaera Syntrophaceticus Pedosphaera Iamia Aciditerrimonas Actinomadura Phormidium Acidopila Candidatus Koribacter Telmatobacter Rhodopila Actinoallomurus Burkholderia Methylosinus Acidobacterium Granulicella Pseudomonas Candidatus Solibacter Erwinia Conexibacter Bryobacter Paenibacillus
Novosphingobium Mycobacterium Luteibacter Ideonella Ewingella Azospirillum Bradyrhizobium Mucilaginibacter Janthinobacterium +
Lysobacter Phenylobacterium Chthoniobacter Acidocella Terriglobus +
Variovorax +
Ferruginibacter +
Pedobacter +
Stenotrophomonas
Rhizobium +
Sphingobacterium +
Chitinophaga +
Sphingomonas +
Devosia +
Rhodococcus
Ochrobactrum +
Frondihabitans +
Achromobacter +
Pseudochrobactrum +
Oidiodendron
Penicillium
Russula
Lactarius
Leptodontidium
Mortierella
Cenococcum
Elaphomyces
Mycoleptodiscus
Tomentella
Odonticium
Xerocomus
Pseudaegerita
Cryptococcus
Campylocarpon
Geomyces
Mycena
Cadophora
Crocicreas
Lachnellula
Rinodina
Candida
Phialocephala
Trichosporon
Devriesia
Meliniomyces
Aspergillus
Troposporella
Athelia
Sympodiella
Asterotremella
Cladosporium
Fulvoflamma
Phyllosticta
Cryptosporiopsis
Epicoccum
Dwayaangam
Cylindrosympodium
Pochonia
Pseudeurotium
Fusarium
Gloiodon
Calcarisporium
Mucor
Kappamyces
Verticillium
Monographella
Mycolevis
Pseudeurotium
Mycena
Meliniomyces
Aspergillus
Russula
Penicillium Calcarisporium
Trichosporon
Monographella +
Sympodiella
Xerocomus +
Mucor
Mortierella
Cladosporium +
Cryptococcus +
Epicoccum +
Rinodina
Athelia
Cryptosporiopsis
Dwayaangam
Candida +
Crocicreas +
Leptodontidium
Verticillium
Troposporella +
Fusarium +
Devriesia
Geomyces
Pochonia +
Asterotremella +
Cylindrosympodium
Cadophora +
Oidiodendron
Elaphomyces
Lactarius
Phialocephala +
Tomentella
Lachnellula
Kappamyces +
Odonticium
Phyllosticta
Gloiodon
Mycolevis
Fulvoflamma
Pseudaegerita
Cenococcum
Campylocarpon
Mycoleptodiscus
!
Week 21
Week 15
Week 9
Week 3
Litter
Litter
Week 2
(d)
Soil
Mycobags soil
Litter
(c)
Fungi
Litter
Mucilaginibacter
Burkholderia
Luteibacter
Granulicella
Sphingomonas
Pedobacter
Bradyrhizobium
Phenylobacterium
Ideonella
Lysobacter
Acidobacterium
Ewingella
Novosphingobium
Terriglobus
Chitinophaga
Frondihabitans
Chthoniobacter
Edaphobacter
Variovorax
Rhizobium
Mycobacterium
Ferruginibacter
Acidopila
Frankia
Conexibacter
Methylosinus
Acidisphaera
Janthinobacterium
Sphingobacterium
Candidatus Solibacter
Pseudomonas
Azospirillum
Devosia
Syntrophaceticus
Pedosphaera
Achromobacter
Rhodopila
Acidocella
Telmatobacter
Actinoallomurus
Paenibacillus
Actinomadura
Aciditerrimonas
Bryobacter
Candidatus Koribacter
Iamia
Phormidium
Erwinia
Ochrobactrum
Stenotrophomonas
Rhodococcus
Pseudochrobactrum
Mycobags litter
Telmatobacter
Acidopila
Granulicella
Candidatus Koribacter
Acidobacterium
Iamia
Aciditerrimonas
Candidatus Solibacter
Mucilaginibacter
Pseudomonas
Paenibacillus
Actinoallomurus
Actinomadura
Pedosphaera
Bradyrhizobium
Conexibacter
Bryobacter
Phenylobacterium
Azospirillum
Burkholderia
Phormidium
Rhodopila
Luteibacter
Acidocella
Edaphobacter
Syntrophaceticus
Chitinophaga
Acidisphaera
Methylosinus
Chthoniobacter
Pedobacter
Mycobacterium
Terriglobus
Sphingobacterium
Ferruginibacter
Lysobacter
Frankia
Frondihabitans
Sphingomonas
Variovorax
Ideonella
Devosia
Achromobacter
Rhizobium
Ochrobactrum
Ewingella
Novosphingobium
Janthinobacterium
Pseudochrobactrum
Stenotrophomonas
Erwinia
Rhodococcus
Week 2
(b)
Soil
Mycobags soil
Bacteria
Litter
(a)
Mycobags litter
1376 Research
Soil
Mortierella Cadophora
Aspergillus Oidiodendron +
Cryptococcus +
Leptodontidium +
Penicillium Epicoccum Russula +
Elaphomyces
Pseudeurotium +
Cladosporium +
Fulvoflamma
Meliniomyces
Cylindrosympodium
Calcarisporium +
Asterotremella +
Mucor +
Dwayaangam
Pseudaegerita
Geomyces +
Fusarium +
Verticillium +
Kappamyces +
Pochonia +
Trichosporon +
Phyllosticta
Sympodiella
Crocicreas +
Rinodina
Candida
Devriesia
Lactarius +
Xerocomus +
Lachnellula
Troposporella
Monographella +
Tomentella +
Phialocephala +
Campylocarpon +
Mycena
Athelia +
Mycoleptodiscus
Cryptosporiopsis
Cenococcum
Odonticium
Gloiodon
Relative abundance (%)
Fig. 5 Abundance of bacterial and fungal genera associated with decomposing fungal mycelia of Tylopilus felleus in forest litter and soil. (a, c) Comparison
of the mean abundance of bacterial and fungal genera in bulk litter and soil and on decomposing fungal mycelia in the corresponding horizon; asterisks
denote taxa with significantly higher abundance on fungal mycelia; lines connect the same genera abundant on fungal mycelia. (b, d) Abundance of
bacterial and fungal genera in the mycobags over time; + and – indicate a significant increase and decrease in abundance with time, respectively. Coloured
dots indicate the known ecophysiology of fungal genera: red, parasite/pathogen (!, known mycoparasite); blue, ectomycorrhizal root symbiont; green,
saprotroph.
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phases of mycelium decomposition were distinguishable in the
mycobags incubated in the soil, the development of fungal communities over time in the mycobags incubated in the litter was
less distinct (Fig. 3).
The background fungal community in the soil was rich in the
sequences of the orders Eurotiales, Russulales and Helotiales,
whereas Helotiales, Corticiales and Agaricales were most abundant
in the litter (Fig. 4). In the mycobags incubated in the soil, most
of these dominant orders were absent, and the community was
mostly dominated by Mortierellales (53%), and further by other
Leotimycetes and Eurotiales. In the mycobags incubated in the litter, Eurotiales (50%) and Capnoidales (30%) were highly abundant (Fig. 4).
As in bacteria, many common fungal taxa from the surrounding soil and the litter, for example, Sympodiella, Odonticium and
ECM fungal genera, were absent from the mycobags, or their
abundance was very low (Fig. 5; Table S4). The mycobags were
specifically enriched in the sequences of Penicillium, Aspergillus,
Crocicreas, Tomentella, Epicoccum, Mucor, Candida and
Verticillium when decomposed in the litter and in Mortierella,
Aspergillus, Cladosporium, Mucor, and Kappamyces when decomposed in the soil. Sequences of the genera Penicillium,
Cladosporium, Aspergillus and Mortierella were highly abundant
in the mycobags, often in tens of per cent and up to 87% (Fig. 5).
The fungal communities in the early stages of mycelial decomposition were almost exclusively composed of saprotrophs, whereas
the parasitic and ECM fungi increased later.
ECM fungi were frequently detected only in soil (36%),
whereas their abundance in litter (3%) and the mycobags (< 5–
8%) was low (Fig. 5). The abundance of the ECM fungi in the
mycobags incubated in the soil was significantly lower than that
in the soil itself (P = 0.0007). Compared with the surrounding
soil and litter, the fungal communities in the mycobags had significantly lower diversity (P = 0.0019). The OTU richness at
1000 sequences/sample was on average 195 8 in control litter
and 108 8 in soil but was as little as 12–70 in the mycobags
incubated in the litter and 60–100 in the mycobags incubated in
the soil; the diversity increased with time. The diversity and evenness in the mycobags incubated in the soil at the end of the experiment were comparable to those of the surrounding soil but not
to those of the litter, where they were clearly reduced.
Discussion
Decomposition of fungal mycelia
In this study, rapid initial decomposition of dead fungal
mycelium was observed, with a loss of 48% of the dry mass within
the first 3 wk. This result is in agreement with those of previous
studies that also observed the rapid initial decomposition of fungal mycelia (Koide & Malcolm, 2009; Drigo et al., 2012; Fernandez & Koide, 2012, 2014; Zeglin & Myrold, 2013). The lower
initial decomposition rate observed in the litter may be
attributable to either a higher fluctuation in abiotic factors such as
temperature and moisture compared with soil or a negative feedback effect of the nutrient-rich litter (Olander & Vitousek, 2000).
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The high initial rates of decomposition are in contrast with
those observed in the tree litter at the same site, where a loss of
only 16% mass was achieved after 4 months (Snajdr et al., 2011).
The decomposition of the plant litter, however, continued
steadily, with 68% of the mass lost in 2 yr (Snajdr et al., 2011).
Although they were low, the decomposition rates of the plant litter in the second year of decomposition were comparable to or
even higher than those for the fungal mycelium at the end of the
21-wk experiment. The observed slow decomposition and the
finding that a similar remaining mycelial mass of 25% was left in
both horizons may indicate that this mycelium fraction is recalcitrant. Partly decomposed rhizomorphs are known to persist over
long periods in forest soils (Clemmensen et al., 2013). Ericoid
mycorrhizal ascomycetes dominating deeper soil horizons of
boreal forests produced melanized hyphae resistant to decomposition and were proposed to potentially facilitate long-term
humus build-up in boreal forests. The presence of ericoid mycorrhizal fungi with melanized hyphae may thus be one reason why
deeper soil horizons are important C sinks (Clemmensen et al.,
2013, 2015; Ekblad et al., 2013). In this study, we have shown
that even the mycelia of common ECM fungi without apparent
melanization contain a significant proportion of recalcitrant matter, which probably contributes to humus formation.
The decomposing fungal mycelium exhibited high enzyme
activity, partly as a consequence of enzymes that were produced
by T. felleus before mycelial decomposition began and which also
probably contributed to the activity observed later. This high
enzyme activity also probably contributed to the rapid initial
decomposition and is consistent with the observations of elevated
chitinase activity in soils enriched with fungal biomass (Zeglin &
Myrold, 2013; Zhao et al., 2013). The range of enzymes that had
high activity in decomposing mycelia was wide and covered
enzymes that decompose various polysaccharides, proteins and
lipids and act in phosphate acquisition. Despite the observed
high enzyme activities at the end of the experiment, the decomposition virtually stopped, which might indicate that the chemical composition of this residual matrix was already highly
transformed and that the relevant substrates were no longer present. The C : N ratio in the decomposing mycelia declined with
time. It was recently proposed that organic N can be translocated
by ECM fungi and saprotrophic cord-forming fungi from N-rich
patches to N-depleted patches to assist decomposition (Boberg
et al., 2014). In this study, N tended to remain in the substrate,
which is consistent with the observation that bacteria and
micromycelial fungi were most abundant while cord-forming
saprotrophs and ECM fungi were scarce for the period of the
experiment; their presence and N translocation, however, cannot
be excluded in later phases of decomposition.
The decomposing mycelium was also demonstrated to be associated with high bacterial biomass but, interestingly, not with
high fungal biomass (Fig. 1). This finding indicates that
mycelium decomposition supports the formation of bacterial
biomass and that bacteria may be more important in its decomposition. Those bacterial genera that were found in association
with decomposition represent a higher share of the total bacterial
community in soil than in litter (22% and 12%, respectively),
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which indicates the greater importance of dead fungal biomass as
a nutrient source in soil than in the litter horizon, with plant litter as the main nutrient source (Snajdr et al., 2008).
Bacteria associated with decomposing fungal mycelia
Our results show that decomposing mycelia are inhabited by a
specific community of bacteria; this is similar to the rhizosphere
and ectomycorrhizosphere, which also harbour unique bacterial
communities (Uroz et al., 2010; Vik et al., 2013). Interestingly,
although the pH of the decomposing mycelia differed significantly in litter and soil, they harboured similar bacterial communities. Instead, the composition of the bacterial community
differed between the early phase of rapid decomposition and the
late phase. Especially in the early phase, the community showed
limited diversity and was dominated by Pseudomonas and
Ewingella, whereas Pedobacter and a range of other bacteria dominated the late phase (Fig. 5). Both Pedobacter and Pseudomonas
species are generalists and possess a wide array of enzymes that are
capable of degrading a diverse set of C sources (Janssen, 2006;
Gordon et al., 2009). More importantly, members of the genera
Pedobacter, Pseudomonas and Ewingella produce chitinase, which
facilitates access to the N contained in the chitin of fungal cell
walls (Inglis & Peberdy, 1996; De Boer et al., 1998; Nissinen
et al., 2012). Ewingella americana and Pseudomonas are mycoparasites (Inglis & Peberdy, 1996; Chowdhury et al., 2007), and this
is consistent with our observation that they participate in the initial decomposition of the mycelia which probably has a similar
composition to the living mycelium. Chitinophaga, a filamentous
chitinolytic gliding bacterium, was previously reported to use fungal hyphae and insects as sources of chitin (Sangkhobol & Skerman, 1981). This genus is one of the most abundant in several
distinctly different sites in South and North America, where it
can represent 7–14% of all sequences (Fulthorpe et al., 2008).
This finding may indicate the importance of fungal mycelia as a
nutrient source in a wide range of soils. Among other myceliumassociated bacteria, Variovorax is a chitinolytic genus that has
been detected in various environments and is able to degrade
complex organic structures, including xenobiotics (Bers et al.,
2011), and Stenotrophomonas was isolated from fungal hyphae
(Gahan & Schmalenberger, 2014) and has been reported to produce N-acetylglucoseaminidase (Yoon et al., 2006). Pseudomonas,
Variovorax and Stenotrophomonas belong to the oxalotrophic Proteobacteria which are capable of becoming dispersed on fungal
mycelia via the ‘fungal highways’, the mycelial networks that
allow the active movement of bacteria in soil (Sahin, 2003; Bravo
et al., 2013). Their association with decomposing mycelia may
indicate that, in addition to movement, fungal hyphae may serve
as their nutrient source.
Fungi associated with decomposing fungal mycelia
As in bacteria, the fungal community on decomposing mycelia
included specific taxa and was less diverse than in the surrounding soil and litter. Interestingly, and in contrast to bacteria,
mycelia in litter and soil were colonized by different fungal taxa
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(Fig. 4). Because of the filamentous nature of fungi, the potential
of a taxon to colonize a novel substrate is important for successful
establishment and determines the following succession. This pattern, called the ‘priority effect’, is well known from decomposing
wood (Hiscox et al., 2015). One may theorize that the priority
effect could explain why the communities in mycobags in the litter differed from those in the soil if they are preferentially colonized by taxa with high abundances in each horizon. However,
this was not the case because the relative abundances of the three
most important primary colonizers of fungal mycelia,
Penicillium, Mortierella, and Aspergillus, were similar in the two
mycobag types. Thus, the probability of initial colonization was
driven by other factors that affected competition, which might
include moisture content or pH.
As in bacteria, the fungal community composition in the
mycobags differed between the early phase of rapid decomposition and the late phase (Fig. 4). Despite the differences in fungal
community between the mycobags incubated in the litter and the
soil, both passed through a period of limited diversity and evenness. Compared with the bulk soil and litter, the mycobags were
specifically rich in nonbasidiomycetous r-strategist species, with
ECM fungi absent. The communities showed some similarity to
the early fungal communities in soil with severed hyphae and
plant roots (Lindahl et al., 2010). A definite increase in certain
free-living ascomycetes from the order Helotiales and the genera
Capronia, Penicillium and Mortierella was detected only 5 d after
disturbance and indicated their involvement in the opportunistic
decomposition of freshly dead mycelia (Lindahl et al., 2010).
The fast incorporation of mycelial C from dead ECM mycelia
into basidiomycetous fungi observed by (Drigo et al., 2012) was
not detected in our study, as the basidiomycete abundance was
low during the fast initial decomposition.
Decomposing leaf litter was found to be markedly enriched in
the genera Mortierella, Cladosporium and Trichosporon at the
same site (Vorıskova & Baldrian, 2013). Mortierella and
Trichosporon were associated with older litter, whereas
Cladosporium was detected at the beginning of the litter decomposition. Because mycelia of saprotrophic fungi are produced on
decomposing litter, it is possible that these fungi may act as
decomposers of fungal mycelia rather than the litter itself. The
decomposition of chitin is a common trait of most fungi because
this activity is required for the transformation of their own
mycelia (Baldrian et al., 2011; Eichlerova et al., 2015), but the
production of processive endochitinases that can efficiently cleave
extracellular chitin is less common. Penicillium spp., Geomyces sp.
and Cladosporium sp. strains from the study area were tested for
endochitinase production, and only Penicillium produced this
enzyme (Baldrian et al., 2011). The latter two fungi might have
used other hydrolytic enzymes, such as a- and b-galactosidases
and glucosidases, to decompose other constituents of the fungal
cell walls (Baldrian et al., 2011). Mycelium decomposition does
not seem to be the only option for these taxa because all of them
were able to degrade cellulose with a processive exocellulase
(Baldrian et al., 2011).
We conclude that dead fungal mycelia represent a unique substrate in the forest ecosystem that is N-rich and contains easily
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decomposable and recalcitrant compounds. Especially in soil,
patches of decomposing fungal mycelia may represent hotspots
of decomposition and bacterial abundance and host a specific
community of bacteria and fungi that successively changes with
the ongoing transformation of the original substrate. Although
our results indicate that the role of bacteria in decomposition of
T. felleus mycelium is especially important, further research is
needed to link individual taxa to the utilization of specific compounds by using genome sequencing, metagenomics or metatranscriptomics. Additionally, the further fate of the recalcitrant
fraction of the mycelia requires more attention in the future
because of its potentially important role in C sequestration in
forest soils. It also must be noted that this study, which only used
mycelia from one fungal species, should be extended by using the
mycelia of other fungi before coming to general conclusions.
Acknowledgements
This work was supported by the Czech Science Foundation (504/
12/P107) and by the research concept of the Institute of
Microbiology of the ASCR (RVO61388971).
Author contributions
V.B. designed the experiment, collected samples, performed the
chemical analyses of the samples and the analyses of microbial
community composition, and wrote the draft of the paper. A.D.
and M.N. contributed to sample collection and performed
microbial community end enzyme analyses. P.B. contributed to
the experimental design and manuscript preparation.
References
Amann RI, Ludwig W, Schleifer KH. 1995. Phylogenetic identification and insitu detection of individual microbial cells without cultivation. Microbiological
Reviews 59: 143–169.
Baldrian P, Kolarık M, Stursova M, Kopecky J, Valaskova V, Vetrovsky T,
ca kova L, Zif
Snajdr J, Rıdl J, Vlcek C et al. 2012. Active and total microbial
communities in forest soil are largely different and highly stratified during
decomposition. ISME Journal 6: 248–258.
Baldrian P, Merhautova V, Cajthaml T, Petra nkova M, Snajdr J. 2010. Smallscale distribution of extracellular enzymes, fungal, and bacterial biomass in
Quercus petraea forest topsoil. Biology and Fertility of Soils 46: 717–726.
Baldrian P, Snajdr J, Merhautova V, Dobiasova P, Cajthaml T, Valaskova V.
2013a. Responses of the extracellular enzyme activities in hardwood forest to
soil temperature and seasonality and the potential effects of climate change. Soil
Biology & Biochemistry 56: 60–68.
Baldrian P, Valaskova V. 2008. Degradation of cellulose by basidiomycetous
fungi. FEMS Microbiology Reviews 32: 501–521.
Baldrian P, Vetrovsk
y T, Cajthaml T, Dobiasova P, Petrankova M, Snajdr J,
Eichlerova I. 2013b. Estimation of fungal biomass in forest litter and soil.
Fungal Ecology 6: 1–11.
Baldrian P, Vorıskova J, Dobiasova P, Merhautova V, Lisa L, Valaskova V.
2011. Production of extracellular enzymes and degradation of biopolymers by
saprotrophic microfungi from the upper layers of forest soil. Plant and Soil
338: 111–125.
Bartnicki-Garcia S. 1968. Cell wall chemistry, morphogenesis and taxonomy of
fungi. Annual Reviews of Microbiology 22: 87–108.
Berg B. 2000. Litter decomposition and organic matter turnover in northern
forest soils. Forest Ecology and Management 133: 13–22.
Ó 2016 The Authors
New Phytologist Ó 2016 New Phytologist Trust
Research 1379
Berlemont R, Martiny AC. 2015. Genomic potential for polysaccharide
deconstruction in bacteria. Applied and Environmental Microbiology 81: 1513–
1519.
Bers K, Sniegowski K, Albers P, Breugelmans P, Hendrickx L, De Mot R,
Springael D. 2011. A molecular toolbox to estimate the number and diversity
of Variovorax in the environment: application in soils treated with the
phenylurea herbicide linuron. FEMS Microbiology Ecology 76: 14–25.
Boberg JB, Finlay RD, Stenlid J, Ekblad A, Lindahl BD. 2014. Nitrogen and
carbon reallocation in fungal mycelia during decomposition of boreal forest
litter. PLoS ONE 9: e92897.
de Boer W, Folman LB, Summerbell RC, Boddy L. 2005. Living in a fungal
world: impact of fungi on soil bacterial niche development. FEMS Microbiology
Reviews 29: 795–811.
Bostrom B, Comstedt D, Ekblad A. 2007. Isotope fractionation and 13C
enrichment in soil profiles during the decomposition of soil organic matter.
Oecologia 153: 89–98.
Bravo D, Cailleau G, Bindschedler S, Simon A, Job D, Verrecchia E, Junier P.
2013. Isolation of oxalotrophic bacteria able to disperse on fungal mycelium.
FEMS Microbiology Letters 348: 157–166.
Bremner JM. 1960. Determination of nitrogen in soil by the Kjeldahl method.
Journal of Agricultural Science 55: 11–33.
Cairney JWG. 2012. Extramatrical mycelia of ectomycorrhizal fungi as
moderators of carbon dynamics in forest soil. Soil Biology and Biochemistry 47:
198–208.
Cao RH, Liu XG, Gao KX, Mendgen K, Kang ZS, Gao JF, Dai Y, Wang X.
2009. Mycoparasitism of endophytic fungi isolated from reed on soilborne
phytopathogenic fungi and production of cell wall-degrading enzymes in vitro.
Current Microbiology 59: 584–592.
Chowdhury PR, Pay J, Braithwaite M. 2007. Isolation, identification and
ecology of Ewingella americana (the causal agent of internal stipe necrosis) from
cultivated mushrooms in New Zealand. Australasian Plant Pathology 36: 424–
428.
Clemmensen KE, Bahr A, Ovaskainen O, Dahlberg A, Ekblad A, Wallander H,
Stenlid J, Finlay RD, Wardle DA, Lindahl BD. 2013. Roots and associated
fungi drive long-term carbon sequestration in boreal forest. Science 339: 1615–
1618.
Clemmensen KE, Finlay RD, Dahlberg A, Stenlid J, Wardle DA, Lindahl BD.
2015. Carbon sequestration is related to mycorrhizal fungal community shifts
during long-term succession in boreal forests. New Phytologist 205: 1525–1536.
Cole JR, Wang Q, Cardenas E, Fish J, Chai B, Farris RJ, Kulam-SyedMohideen AS, McGarrell DM, Marsh T, Garrity GM et al. 2009. The
Ribosomal Database Project: improved alignments and new tools for rRNA
analysis. Nucleic Acids Research 37: D141–D145.
Colpaert JV, VanLaere A, VanAssche JA. 1996. Carbon and nitrogen allocation
in ectomycorrhizal and non-mycorrhizal Pinus sylvestris L. seedlings. Tree
Physiology 16: 787–793.
Cooke RC, Whipps JM. 1993. Ecophysiology of fungi. Oxford, UK: WileyBlackwell.
De Boer W, Gunnewiek P, Lafeber P, Janse JD, Spit BE, Woldendorp JW.
1998. Anti-fungal properties of chitinolytic dune soil bacteria. Soil Biology &
Biochemistry 30: 193–203.
Dowd SE, Callaway TR, Wolcott RD, Sun Y, McKeehan T, Hagevoort RG,
Edrington TS. 2008. Evaluation of the bacterial diversity in the feces of cattle
using 16S rDNA bacterial tag-encoded FLX amplicon pyrosequencing
(bTEFAP). BMC Microbiology 8: 125.
Drigo B, Anderson IC, Kannangara GSK, Cairney JWG, Johnson D. 2012.
Rapid incorporation of carbon from ectomycorrhizal mycelial necromass into
soil fungal communities. Soil Biology & Biochemistry 49: 4–10.
Edgar RC. 2010. Search and clustering orders of magnitude faster than BLAST.
Bioinformatics 26: 2460–2461.
Edgar RC, Haas BJ, Clemente JC, Quince C, Knight R. 2011. UCHIME improves
sensitivity and speed of chimera detection. Bioinformatics 27: 2194–2200.
ca kova L, Lisa L, Dobiasova P, Baldrian P. 2015.
Eichlerova I, Homolka L, Zif
Enzymatic systems involved in decomposition reflects the ecology and
taxonomy of saprotrophic fungi. Fungal Ecology 13: 10–22.
Ekblad A, Wallander H, Godbold DL, Cruz C, Johnson D, Baldrian P, Bjork
RG, Epron D, Kieliszewska-Rokicka B, Kjoller R et al. 2013. The production
New Phytologist (2016) 210: 1369–1381
www.newphytologist.com
1380 Research
and turnover of extramatrical mycelium of ectomycorrhizal fungi in forest soils:
role in carbon cycling. Plant and Soil 366: 1–27.
Fernandez CW, Koide RT. 2012. The role of chitin in the decomposition of
ectomycorrhizal fungal litter. Ecology 93: 24–28.
Fernandez CW, Koide RT. 2014. Initial melanin and nitrogen concentrations
control the decomposition of ectomycorrhizal fungal litter. Soil Biology &
Biochemistry 77: 150–157.
Fulthorpe RR, Roesch LFW, Riva A, Triplett EW. 2008. Distantly sampled soils
carry few species in common. ISME Journal 2: 901–910.
Gahan J, Schmalenberger A. 2014. The role of bacteria and mycorrhiza in plant
sulfur supply. Frontiers in Plant Sciences 5: 723.
Gordon NS, Valenzuela A, Adams SM, Ramsey PW, Pollock JL, Holben WE,
Gannon JE. 2009. Pedobacter nyackensis sp nov., Pedobacter alluvionis sp nov
and Pedobacter borealis sp nov., isolated from Montana flood-plain sediment
and forest soil. International Journal of Systematic and Evolutionary Microbiology
59: 1720–1726.
Guthrie JL, Castle AJ. 2006. Chitinase production during interaction of
Trichoderma aggressivum and Agaricus bisporus. Canadian Journal of
Microbiology 52: 961–967.
van der Heijden MGA, Martin FM, Selosse M-A, Sanders IR. 2015.
Mycorrhizal ecology and evolution: the past, the present, and the future. New
Phytologist 205: 1406–1423.
Hendricks JJ, Mitchell RJ, Kuehn KA, Pecot SD. 2016. Ectomycorrhizal
fungal mycelia turnover in a longleaf pine forest. New Phytologist 209: 1693–
1704.
Ihrmark K, Asmail N, Ubhayasekera W, Melin P, Stenlid J, Karlsson M. 2010.
Comparative molecular evolution of Trichoderma chitinases in response to
mycoparasitic interactions. Evolutionary Bioinformatics 6: 1–25.
Inglis PW, Peberdy JF. 1996. Isolation of Ewingella americana from the
cultivated mushroom, Agaricus bisporus. Current Microbiology 33: 334–337.
Janssen PH. 2006. Identifying the dominant soil bacterial taxa in libraries of 16S
rRNA and 16S rRNA genes. Applied and Environmental Microbiology 72:
1719–1728.
Koide RT, Malcolm GM. 2009. N concentration controls decomposition rates of
different strains of ectomycorrhizal fungi. Fungal Ecology 2: 197–202.
Koljalg U, Nilsson RH, Abarenkov K, Tedersoo L, Taylor AFS, Bahram M,
Bates ST, Bruns TD, Bengtsson-Palme J, Callaghan TM et al. 2013. Towards
a unified paradigm for sequence-based identification of fungi. Molecular Ecology
22: 5271–5277.
Leveau JHJ, Preston GM. 2008. Bacterial mycophagy: definition and diagnosis
of a unique bacterial–fungal interaction. New Phytologist 177: 859–876.
Lindahl BD, de Boer W, Finlay RD. 2010. Disruption of root carbon transport
into forest humus stimulates fungal opportunists at the expense of mycorrhizal
fungi. ISME Journal 11: 872–881.
Lindahl BD, Ihrmark K, Boberg J, Trumbore SE, H€ogberg P, Stenlid J, Finlay
RD. 2007. Spatial separation of litter decomposition and mycorrhizal nitrogen
uptake in a boreal forest. New Phytologist 173: 611–620.
Lindahl BD, Nilsson RH, Tedersoo L, Abarenkov K, Carlsen T, Kjoller R,
Koljalg U, Pennanen T, Rosendahl S, Stenlid J et al. 2013. Fungal
community analysis by high-throughput sequencing of amplified markers – a
user’s guide. New Phytologist 199: 288–299.
Lopez-Mondejar R, Vorıskova J, Vetrovsky T, Baldrian P. 2015. The
bacterial community inhabiting temperate deciduous forests is vertically
stratified and undergoes seasonal dynamics. Soil Biology & Biochemistry 87:
43–50.
Lundberg DS, Lebeis SL, Paredes SH, Yourstone S, Gehring J, Malfatti S,
Tremblay J, Engelbrektson A, Kunin V, del Rio TG et al. 2012.
Defining the core Arabidopsis thaliana root microbiome. Nature 488:
86–90.
Majdi H, Truus L, Johansson U, Nylund JE, Wallander H. 2008. Effects of
slash retention and wood ash addition on fine root biomass and production and
fungal mycelium in a Norway spruce stand in SW Sweden. Forest Ecology and
Management 255: 2109–2117.
Meyer F, Paarmann D, D’Souza M, Olson R, Glass EM, Kubal M, Paczian T,
Rodriguez A, Stevens R, Wilke A et al. 2008. The metagenomics RAST server
– a public resource for the automatic phylogenetic and functional analysis of
metagenomes. BMC Bioinformatics 9: 386.
New Phytologist (2016) 210: 1369–1381
www.newphytologist.com
New
Phytologist
Mouginot C, Kawamura R, Matulich KL, Berlemont R, Allison SD, Amend AS,
Martiny AC. 2014. Elemental stoichiometry of Fungi and Bacteria strains from
grassland leaf litter. Soil Biology & Biochemistry 76: 278–285.
Nilsson K, Bjurman J. 1998. Chitin as an indicator of the biomass of two wooddecay fungi in relation to temperature, incubation time, and media
composition. Canadian Journal of Microbiology 44: 575–581.
Nissinen RM, Mannisto MK, van Elsas JD. 2012. Endophytic bacterial
communities in three arctic plants from low arctic fell tundra are cold-adapted
and host-plant specific. FEMS Microbiology Ecology 82: 510–522.
Olander LP, Vitousek PM. 2000. Regulation of soil phosphatase and chitinase
activity by N and P availability. Biogeochemistry 49: 175–190.
Osono T. 2007. Ecology of ligninolytic fungi associated with leaf litter
decomposition. Ecological Research 22: 955–974.
Osono T, Takeda H. 2005. Decomposition of organic chemical components in
relation to nitrogen dynamics in leaf litter of 14 tree species in a cool temperate
forest. Ecological Research 20: 41–49.
Prevost-Boure N, Chemidin N, Christen R, Dequiedt S, Mougel C, Lelievre M,
Jolivet C, Shahbazkia HR, Guillou L, Arrouays D et al. 2011. Validation and
application of a PCR primer set to quantify fungal communities in the soil
environment by real-time quantitative PCR. PLoS ONE 6: e24166.
Reeder J, Knight R. 2010. Rapidly denoising pyrosequencing amplicon reads by
exploiting rank-abundance distributions. Nature Methods 7: 668–669.
Sa gova-Mareckova M, Cerm
a k L, Novotna J, Plhackova K, Forstova J, Kopecky
J. 2008. Innovative methods for soil DNA purification tested in soils with
widely differing characteristics. Applied and Environmental Microbiology 74:
2902–2907.
Sahin N. 2003. Oxalotrophic bacteria. Research in Microbiology 154: 399–407.
Sangkhobol V, Skerman VBD. 1981. Chitinophaga, a new genus of chitinolytic
myxobacteria. International Journal of Systematic Bacteriology 31: 285–293.
Seidl V. 2008. Chitinases of filamentous fungi: a large group of diverse proteins
with multiple physiological functions. Fungal Biology Reviews 22: 36–42.
Snajdr J, Cajthaml T, Valaskova V, Merhautova V, Petra nkova M, Spetz P,
Leppanen K, Baldrian P. 2011. Transformation of Quercus petraea litter:
successive changes in litter chemistry are reflected in differential enzyme activity
and changes in the microbial community composition. FEMS Microbiology
Ecology 75: 291–303.
Snajdr J, Valaskova V, Merhautova V, Herinkova J, Cajthaml T, Baldrian P.
2008. Spatial variability of enzyme activities and microbial biomass in the
upper layers of Quercus petraea forest soil. Soil Biology & Biochemistry 40:
2068–2075.
Soudzilovskaia NA, van der Heijden MGA, Cornelissen JHC, Makarov MI,
Onipchenko VG, Maskeri B, Maslov MN, Akhmetzhanova AA, van Bodegom
PM. 2015. Quantitative assessment of the differential impacts of arbuscular
and ectomycorrhiza on soil carbon cycling. New Phytologist 208: 280–293.
Stursova M, Baldrian P. 2011. Effects of soil properties and management on the
activity of soil organic matter transforming enzymes and the quantification of
soil-bound and free activity. Plant and Soil 338: 99–110.
Stursova M, Zifca kova L, Leigh MB, Burgess R, Baldrian P. 2012. Cellulose
utilization in forest litter and soil: identification of bacterial and fungal
decomposers. FEMS Microbiology Ecology 80: 735–746.
Tedersoo L, Bahram M, Polme S, Koljalg U, Yorou NS, Wijesundera R,
Villarreal Ruiz L, Vasco-Palacios AM, Thu PQ, Suija A et al. 2014. Fungal
biogeography. Global diversity and geography of soil fungi. Science 346:
1256688.
Uroz S, Buee M, Murat C, Frey-Klett P, Martin F. 2010. Pyrosequencing reveals
a contrasted bacterial diversity between oak rhizosphere and surrounding soil.
Environmental Microbiology Reports 2: 281–288.
Vainio EJ, Hantula J. 2000. Direct analysis of wood-inhabiting fungi using
denaturing gradient gel electrophoresis of amplified ribosomal DNA.
Mycological Research 104: 927–936.
Valaskova V, Snajdr J, Bittner B, Cajthaml T, Merhautova V, Hoffichter M,
Baldrian P. 2007. Production of lignocellulose-degrading enzymes and
degradation of leaf litter by saprotrophic basidiomycetes isolated from a
Quercus petraea forest. Soil Biology & Biochemistry 39: 2651–2660.
Vetrovsky T, Baldrian P. 2013. Analysis of soil fungal communities by amplicon
pyrosequencing: current approaches to data analysis and the introduction of the
pipeline SEED. Biology and Fertility of Soils 49: 1027–1037.
Ó 2016 The Authors
New Phytologist Ó 2016 New Phytologist Trust
New
Phytologist
Vetrovsk
y T, Baldrian P, Gabriel J. 2013. Extracellular enzymes of the white-rot
fungus Fomes fomentarius and purification of 1,4-b-glucosidase. Applied
Biochemistry and Biotechnology 169: 100–109.
Vik U, Logares R, Blaalid R, Halvorsen R, Carlsen T, Bakke I, Kolsto AB,
Okstad OA, Kauserud H. 2013. Different bacterial communities in
ectomycorrhizae and surrounding soil. Scientific Reports 3: 3471.
Vorıskova J, Baldrian P. 2013. Fungal community on decomposing leaf litter
undergoes rapid successional changes. ISME Journal 7: 477–486.
Vorıskova J, Brabcova V, Cajthaml T, Baldrian P. 2014. Seasonal dynamics of
fungal communities in a temperate oak forest soil. New Phytologist 201: 269–
278.
Wallander H, Goransson H, Rosengren U. 2004. Production, standing biomass
and natural abundance of 15N and 13C in ectomycorrhizal mycelia collected at
different soil depths in two forest types. Oecologia 139: 89–97.
White TJ, Bruns TD, Lee SB, Taylor JW. 1990. Analysis of phylogenetic
relationship by amplification and direct sequencing of ribosomal RNA genes.
In: Innis MA, Gelfand DH, Sninsky JN, White TJ, eds. PCR protocols: a guide
to methods and applications. New York, NY, USA: Academic press, 315–322.
Wilmotte A, Van der Auwera G, De Wachter R. 1993. Structure of the 16 S
ribosomal RNA of the thermophilic cyanobacterium Chlorogloeopsis HTF
(‘Mastigocladus laminosus HTF’) strain PCC7518, and phylogenetic analysis.
FEBS Letters 317: 96–100.
Yoon JH, Kang SJ, Oh HW, Oh TK. 2006. Stenotrophomonas dokdonensis sp
nov., isolated from soil. International Journal of Systematic and Evolutionary
Microbiology 56: 1363–1367.
Zeglin LH, Myrold DD. 2013. Fate of decomposed fungal cell wall material in
organic horizons of old-growth Douglas-fir forest soils. Soil Science Society of
America Journal 77: 489–500.
Zhao J, Xue QH, Niu GG, Xue L, Shen GH, Du JZ. 2013. Extracellular enzyme
production and fungal mycelia degradation of antagonistic Streptomyces
induced by fungal mycelia preparation of cucurbit plant pathogens. Annals of
Microbiology 63: 809–812.
Research 1381
Supporting Information
Additional supporting information may be found in the online
version of this article.
Table S1 Bacterial OTUs associated with the decomposing fungal mycelia of Tylopilus felleus and in the surrounding forest litter
and soil
Table S2 Bacterial genera associated with the decomposing fungal mycelia of Tylopilus felleus and in the surrounding forest litter
and soil
Table S3 Fungal OTUs associated with the decomposing fungal
mycelia of Tylopilus felleus and in the surrounding forest litter
and soil
Table S4 Fungal genera associated with the decomposing fungal
mycelia of Tylopilus felleus and in the surrounding forest litter
and soil
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