Proteome analysis of fungal and bacterial involvement in leaf litter decomposition

RESEARCH ARTICLE

Proteome analysis of fungal and bacterial involvement in

leaf litter decomposition

Thomas Schneider1, Bertran Gerrits2, Regula Gassmann1, Emanuel Schmid1,Mark O. Gessner3, Andreas Richter4, Tom Battin5,6, Leo Eberl1 and Kathrin Riedel1

1 Institute of Plant Biology, Department of Microbiology, University of Zurich, Zurich, Switzerland2 Functional Genomics Center, University and ETH Zurich, Zurich, Switzerland3 Department of Aquatic Ecology Eawag: Swiss Federal Institute of Aquatic Science & Technology,

and Institute of Integrative Biology (IBZ), ETH Zurich, D .ubendorf, Switzerland4 Chemical Ecology and Ecosystem Research, University of Vienna, Vienna, Austria5 Department of Freshwater Ecology, University of Vienna, Vienna, Austria6 Wasser Cluster Lunz, Lunz am See, Austria

Received: October 8, 2009

Revised: January 29, 2010

Accepted: February 4, 2010

Fungi and bacteria are key players in the decomposition of leaf litter, but their individual

contributions to the process and their interactions are still poorly known. We combined semi-

quantitative proteome analyses (1-D PAGE-LC-MS/MS) with qualitative and quantitative

analyses of extracellular degradative enzyme activities to unravel the respective roles of a fungus

and a bacterium during litter decomposition. Two model organisms, a mesophilic Gram-

negative bacterium (Pectobacterium carotovorum) and an ascomycete (Aspergillus nidulans), were

grown in both, pure culture and co-culture on minimal medium containing either glucose or

beech leaf litter as sole carbon source. P. carotovorum grew best in co-culture with the fungus,

whereas growth of A. nidulans was significantly reduced when the bacterium was present. This

observation suggests that P. carotovorum has only limited capabilities to degrade leaf litter and

profits from the degradation products of A. nidulans at the expense of fungal growth. In

accordance with this interpretation, our proteome analysis revealed that most of the

extracellular biodegradative enzymes (i.e. proteases, pectinases, and cellulases) in the cultures

with beech litter were expressed by the fungus, the bacterium producing only low levels of

pectinases.

Keywords:

Bacteria / Degradative enzymes / Fungi / Leaf litter decomposition / Microbiology /

Semi-quantitative proteomics

1 Introduction

Leaf litter decomposition is a key ecosystem process that

greatly contributes to the fluxes of matter and energy in the

biosphere. Leaf litter consists mainly of polymers such as

cellulose, the most abundant carbohydrate on earth, but also

lignin, hemicellulose, pectin, and protein [1, 2], which all

together require numerous enzymes for degradation. Fungi

and bacteria are key to this process by expressing a suite of

extracellular enzymes. In fact, these microorganisms

decompose almost 90% of the plant biomass produced in

terrestrial ecosystems [3].

Fungi are able to degrade even highly recalcitrant litter

[4–6]. Bacteria also produce numerous cell-wall degrading

enzymes, but, compared with fungi, they appear to be

inferior decomposers of leaf litter in streams [7–9]. While

the contribution of fungi and bacteria to plant litter polymer

degradation is generally acknowledged, our understanding

of the underlying molecular mechanisms remains poor.Abbreviation: CFU, colony-forming unit

Correspondence: Dr. Thomas Schneider, Institute of Plant Biol-

ogy, Department of Microbiology, University of Zurich, Winter-

thurerstrasse 190, 8057 Zurich, Switzerland

E-mail: [email protected]

Fax: 141-44-63-50384

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com

Proteomics 2010, 10, 1819–1830 1819DOI 10.1002/pmic.200900691

Analyses of origin, function, and activity of microbial

enzymes can provide new insights into ecosystem processes

[10]: Enzymes have intrinsic biological functions and many

are involved in physiological processes directly linked to

organic matter transformation. However, the multitude of

proteins expressed in natural communities is often a major

obstacle to relating enzyme data to ecosystem-level proces-

ses. One way to overcome this limitation is the application of

proteomics to environmental samples. Indeed, recent large-

scale characterizations of the entire protein complement of

diverse microbial communities have proven useful to

describe microbial functions in activated sludge [11, 12], soil

[13], and lake and ground water [13–15]. In addition,

proteomics can be used to infer the identity of the active

organisms even in multispecies communities. However, a

major limitation of such meta-proteomic approaches to date

is the enormous complexity of natural environments and

the lack of protein sequence information for most

microbiota.

The aim of this study was to assess the contribution of

fungi and bacteria to leaf litter decomposition by combining

measurements of enzyme activities with protein expression

data from semi-quantitative community proteomics. To

circumvent the problem of insufficient sequence informa-

tion, we used a model system consisting of a well-char-

acterized fungus and bacterium. The degradation of major

leaf litter polymers was assessed in both, pure cultures and

co-cultures of the two microorganisms. Pectobacteriumcarotovorum and Aspergillus nidulans were selected as model

organisms because their genomes have been sequenced [16,

17] and their enzymes involved in plant cell-wall degradation

have been characterized in detail [18–20]. We used 1-D

PAGE-LC-MS/MS to characterize the proteome. Proteomics

data were combined with qualitative and quantitative

analyses of the activity of protein-, pectin- and cellulose-

degrading enzymes, which allowed us to attribute the

measured enzyme activities to the respective source organ-

isms. Our results showed that A. nidulans is the main

producer of extracellular degradative enzymes, in particular

of proteases, cellulases, and pectinases, independent of

whether the fungus grows in pure culture or in co-culture

with P. carotovorum. The bacterium was ineffective to

produce extracellular degradative enzymes but appears to

profit from the fungus by utilizing its degradation products,

thereby reducing fungal growth.

2 Materials and methods

2.1 Strains and growth conditions

P. carotovorum subsp. carotovorum (American Type Culture

Collection no. 39048) and A. nidulans (Austrian Center of

Biological Resources and Applied Mycology no. MA5366)

were used as bacterial and fungal model organisms.

P. carotovorum and A. nidulans pure cultures as well as

co-cultures containing both organisms were grown in 1 L

Erlenmeyer flasks containing 200 ml AB minimal medium

[21] supplemented with either 0.2% glucose or 1% sterile

beech leaf litter. Beech litter was collected from several

sampling sites in Austria, pooled, and dried at 301C for 24 h.

The dried litter was fragmented (o0.5 cm2), added to the

medium and autoclaved (1201C, 15 min). Autoclaving was

chosen since X-ray sterilization might be ineffective at

inactivating spore-forming organisms in litter. To exclude

effects on growth and expression rates of extracellular

enzymes caused by a nutrient release from beech leaf litter

during autoclaving we compared P. carotovorum cultures

grown on autoclaved or X-ray sterilized litter; in fact, no

differences in bacterial growth (Supporting Information

Fig. 1) and enzyme expression rates were visible when

analysing the cultures with beech litter by 1-D-LC-MS/MS

after 17 days of incubation (data not shown).

P. carotovorum cultures were inoculated with an overnight

culture grown on minimal medium supplemented with

glucose to a final OD600 of 0.05. A. nidulans cultures were

inoculated with six slices of agar (0.2 cm2 each) containing

freshly grown mycelium. Cultures were incubated for 21

days at 281C with vigorous agitation at 225 rpm. Aliquots

(25 mL) of the culture medium were collected after 0, 3, 6, 9,

13, 17, and 21 days of growth. Aliquots were centrifuged

(3000� g, 15 min, 41C) to remove any litter debris and cells,

then filter-sterilized (Millex GP, PES filters, Millipore,

0.22 mm pore size) and frozen (�201C) until further

processing. All growth experiments were performed in two

independent replicates.

2.2 Bacterial and fungal growth

Bacterial growth was assessed by counting the number of

colony-forming units (CFUs) on LB-plates at the beginning

of the experiment as well as after 3, 6, 13, 17, and 21 days.

Possible inhibitory effects of P. carotovorum and A. nidulanswere assessed by growing both organisms together on LB

agar plates; no growth inhibition was observed while

growing the strains on nutrient-rich medium (data not

shown).

Fungal biomass was determined as ergosterol [22] at the

start of the experiment and after 3, 6, 13, and 21 days. The

concentration of ergosterol per gram biomass varies rela-

tively little over fungal growth stages; therefore, it is a well-

suited marker for changes in fungal biomass [22, 23]. To

determine its concentration, cultures containing leaves and

fungal mycelium were harvested, freeze-dried, and weigh-

ted, and ground sub-samples (50 mg) extracted in 10 mL

alkaline methanol with stirring for 30 min at 801C. The

cooled extracts were passed over SPE cartridges (Waters Sep-

Paks, Vac RC, tC18, 500 mg sorbent) [24], no attempt was

made to raise pH after sample loading and ergosterol

elution, because ergosterol degradation in the acid extracts

was not observed. Final purification was achieved on a high-

1820 T. Schneider et al. Proteomics 2010, 10, 1819–1830


pressure liquid chromatograph (Jasco PU-980 pump, AS-950

autosampler, MD 2010 Plus Multi-wavelength Detector;

Tokyo, Japan). The column was a LichroSpher 100 RP-18

column (0.46� 25 cm) (Merck, Darmstadt, Germany). The

injection volume was 20mL. Ergosterol was quantified by

measuring absorbance at 282 nm.

2.3 Carbohydrate analysis

High-pressure liquid chromatography was also used to

quantify carbohydrates in the culture medium. Carbohy-

drates were separated using a CarboPac PA1 anion exchange

column (4� 250 mm, Dionex, Sunnyvale, CA, USA) and a

linear gradient of NaOH and Na-acetate. Specifically, the

conditions were: 0–3 min with 100 mM NaOH, 3–20 min

with a linear gradient from 100 mM NaOH to final

concentrations of 200 mM NaOH and 100 mM Na-acetate,

and 20–22 min with a linear gradient to a final concentration

of 100 mM NaOH. The column was regenerated at 100 mM

NaOH for 18 min. Pulsed amperometric detection (Dionex

ED 50 detector) was used for carbohydrate detection. The

separated carbohydrates were identified and quantified

using authentic myo-inositol, rhamnose, arabinose, xylose,

glucose, galactose, and sucrose standards at a concentration

of 2.5 nmol/10 mL.

2.4 Spectrophotometric enzyme assays

Filter-sterilized culture supernatants were used for quanti-

tative enzyme assays. Protease activity was assayed with a

modified azocasein hydrolysis assay [25]; for details see

Supporting Information. Pectinase and cellulase activity

were assayed according to [26, 27]; for details see Supporting

Information.

2.5 Qualitative enzyme analysis by zymograms

Filter-sterilized culture supernatants were concentrated 50-

fold by ultrafiltration (10 kDa cutoff membrane; Vivaspin

500, Sigma) and separated by SDS-PAGE [28] using 12%

polyacrylamide gels. To visualize protease, cellulase, and

pectinase activity, 0.2% azocasein, 0.1% carboxymethyl-

cellulose, or 0.1% apple pectin were incorporated into the

gel matrix, respectively. After electrophoresis, proteins were

renaturated by washing the gels twice in 50 mM Tris/HCl

(pH 7.0) and 25% isopropanol for 15 min at room

temperature and twice in 50 mM Tris/HCl (pH 7.0) for

15 min at room temperature. After renaturation, zymo-

grams were incubated at 401C for 3 h. Subsequently, gels

were stained to visualize enzyme activities: (i) azocasein

containing gels were washed in 1 M NaOH for 5 min until

protease activity could be detected as colourless bands

against an orange background; (ii) carboxymethyl-cellulose

containing gels were stained with 0.5% Congo red for 5 min

and destained with 1 M NaCl until cellulase activity was

visible as colourless bands against a red background; and

(iii) pectin containing gels were stained with 0.05% Ruthe-

nium red for 10 min and destained with water until pecti-

nase activity could be detected as colourless bands against a

purple background.

2.6 1-D SDS-PAGE coupled to LC-MS/MS

Proteins in the filter-sterilized culture supernatants were

concentrated 50-fold by ultrafiltration (10 kDa cutoff

membrane; Vivaspin 500) and separated by SDS-PAGE [28]

using 12% polyacrylamide gels. Protein lanes were cut in

three slices and gel slices were immediately subjected to in-

gel tryptic digestion [29]. The resulting peptide mixtures

were analysed on a hybrid LTQ-Orbitrap mass spectrometer

(ThermoFischer Scientific, Bremen, Germany) interfaced

with a nanoelectrospray ion source (for details see

Supporting Information).

2.7 Database searches

The MASCOT Search Engine (version no. 2.2.04) was used

for protein database searches. The reference database

contained proteins from the sequenced bacterial and fungal

strains Pectobacterium atrosepticum SCRI1043 [16], Strepto-myces coelicolor A3 (2) [30] A. nidulans FGSC A4 [17], and

Phanerochaete chrysosporium RP78 [31] that are closely rela-

ted to our model organisms, as well as common contami-

nants like keratin and trypsin (total number of database

entries 33 631). MS/MS ion searches were performed with

the following settings: (i) trypsin was chosen as protein-

digesting enzyme and up to two missed cleavages were

tolerated, (ii) carbamidomethylation of cystein was chosen

as fixed modification, and (iii) oxidation of methionine and

formation of pyro-glutamic acid from glutamine and gluta-

mic acid were chosen as variable modifications. Searches

were performed with a parent ion mass tolerance of 5 ppm

and a fragment ion mass tolerance of 0.8 Da.

2.8 Data validation and protein quantification

Scaffold (version Scaffold_2.02.01, Proteome Software,

Portland, OR, USA) was used to validate and quantify MS/

MS-based peptide and protein identifications. Peptide

identifications were accepted if they could be established at

greater than 95% probability as specified by the Peptide

Prophet algorithm [32]. Protein identifications were accep-

ted if they could be established at greater than 99% prob-

ability and contained at least two identified peptides in one

of the analyzed samples. Protein probabilities were assigned

by the Protein Prophet algorithm [33]. Proteins that

Proteomics 2010, 10, 1819–1830 1821


contained similar peptides and could not be differentiated

based on MS/MS analysis alone were grouped to satisfy the

principles of parsimony. To determine the false-discovery

rate, searches were performed against a composite version

of our self-constructed reference database, created by

concatenating the target protein sequences with reversed

sequences (67 262 entries) as described by Elias and Gygi

[34]. The number of assigned reversed hits was multiplied

by 2 and divided by the total number of identified proteins.

We computed a false-discovery rate lower than 1%. Semi-

quantitative analyses of protein abundances were performed

on the basis of the number of unique peptides that were

assigned to a protein by the Scaffold software.

2.9 Further data processing

Proteins with unknown function were annotated by a BLAST

search against the Swiss-prot/Uni-prot database (400 771

entries) using the BLASTP stand-alone BLAST tool (version

no. 2.2.181). To further categorize the identified proteins, the

corresponding EC numbers were assigned by searching the

Uni-prot Knowledgebase database. All proteins were checked

for the presence of a signal peptide needed for secretion to

confirm extracellular localization using SignalP 3.0 [35, 36].

3 Results

3.1 Growth of P. carotovorum and A. nidulans on

beech leaf litter

Consistently higher bacterial cell numbers were detected when

P. carotovorum was grown on leaf litter in co-culture with

A. nidulans (Fig. 1A). Bacteria reached the highest number of

CFUs after 13 days both in pure culture and in co-culture with

A. nidulans. However, P. carotovorum cells remained viable

much longer when co-cultured with the fungus.

The opposite effect was observed for A. nidulans when

grown in the absence and presence of P. carotovorum (Fig.

1B). In the presence of the bacterium, fungal growth was

reduced and no further increase was detected after day 7.

These observations contrast with the results of pure cultures

where fungal biomass was higher and increased continu-

ously until the end of the experiment.

3.2 Degradative extracellular enzymes

Enzyme activities varied strongly among the different

cultures and over time. Protease, cellulase, and pectinase

activity was detected mainly in pure cultures and co-cultures

of A. nidulans grown on beech litter (Figs. 2 and 3). The only

exception was pectinase activity detected in pure cultures of

P. carotovorum grown on litter. Although the strain is also

capable of producing proteases and cellulases, as indicated

by experiments with glucose as carbon source

(Fig. 3), activities of these enzymes were not detected by

either spectrophotometric assays or zymogram analyses.

Protease activity constantly increased over time in pure

cultures of A. nidulans grown on litter and was much lower

in the presence of P. carotovorum. This inhibition was

observed in growth media with both, glucose (data not

shown) and beech litter (Fig. 2A). The results of the spec-

trophotometric protease assays were corroborated by zymo-

gram analyses (Fig. 3A). After 3 days of growth on glucose, a

protease band of approximately 50 kDa was present in the

supernatant of P. carotovorum pure cultures, whereas

multiple bands within a size range of 10–50 kDa were

detected in A. nidulans cultures. In accordance with the

spectrophotometric measurements, the intensity of the

protease bands was reduced in the co-cultures of A. nidulansand P. carotovorum. In cultures with beech litter, protease

activity was only detected in co-cultures and pure cultures of

A. nidulans (Figs. 2A and 3A). Protease band patterns were

similar in these two culture types.

Figure 1. Growth of P. carotovorum and A. nidulans in pure

culture and co-culture in minimal medium with beech litter. (A)

Growth of P. carotovorum was assessed as CFUs on agar plates.

(B) Growth of A. nidulans was determined by quantifying

ergosterol, a biomarker for fungal biomass. Data represent

means of four independent experiments. Error bars indicate

standard deviation.



Cellulase and pectinase activity of A. nidulans in pure

culture increased rapidly during early growth on beech litter

and slowly increased further or reached a plateau after day 6

(Figs. 2B and C). Activities showed similar temporal

patterns and activity levels in co-cultures, indicating that the

respective enzymes in the co-culture were produced by the

fungus only (Fig. 2). This finding is in accordance with

results of the zymogram analyses (Figs. 3B and C), in which

only samples containing proteins of A. nidulans showed

multiple cellulase or pectinase bands. In contrast to protease

activity, cellulase and pectinase activities of the fungus were

not affected by the presence of the bacterium. Pure cultures

of P. carotovorum on litter showed no cellulase activity and

strongly reduced pectinase activity in the spectro-

photometric assays (Figs. 2B and C). In fact, although

pectinase expression reached an early maximum at day 3, it

declined almost to the control level thereafter. No pectinase

bands were detected in the zymograms (Fig. 3C), possibly

because sensitivity of the zymogram analysis is lower than

that of the spectrophotometric enzyme assay.

3.3 Semi-quantitative proteome analysis

It is known that the number of peptides identified by 1-D

PAGE-LC-MS/MS correlate positively with the abundance of

the corresponding protein [37]; therefore peptide counts can

be used as an indicator of the enzyme expression rates of a

respective source organism.

A total of 101 extracellular degradative enzymes were

identified in the secretome of the different cultures

(Supporting Information Table 1). They included protein-,

cellulose-, pectin-, chitin-, xylan-, cutin, lipid-, starch-, and

mannan-degrading enzymes. In line with results from the

spectrophotometric and zymogram analyses, most of the

enzymes in the cultures supplemented with beech litter

were produced by A. nidulans, with P. carotovorum expres-

sing only small amounts of some pectinases (Fig. 4).

3.4 Proteases

No extracellular proteases were identified in the secretome

of P. carotovorum grown on beech litter as sole carbon source

(Figs. 4B and C) and when the bacterium was grown on

glucose, only one metalloprotease (EC 3.4.24.-; gi|50121709)

was identified (Fig. 4A, Supporting Information Table 1). By

contrast, 11 fungal extracellular proteases were identified in

cultures with litter (Figs. 4B and C; Supporting Information

Table 1), production of which increased over time. They

included three putative carboxypeptidases (AN1226,

AN2237, AN7121), two aminopeptidases (EC 3.4.11.15;

AN3918, AN8445), two neutral proteases (EC 3.4.24.39;

AN3959, AN7962), an alkaline protease (EC 3.4.21.-;

AN5558), a putative dipeptidyl aminopeptidase (EC 3.4.14.-;

AN6438), a tripeptidyl peptidase (EC 3.4.14.9; AN7159), and

a putative serine protease (EC 3.4.-.-; AN7231).

3.5 Cellulases

Cellulases were not detected when P. carotovorum was

grown on litter (Figs. 4E and F), but with glucose as carbon

source, endoglucanase V was found (EC 3.2.1.4;

gi|50120908; Fig. 4D, Supporting Information Table 1).

Twenty-one different fungal cellulose-degrading proteins

were identified in litter cultures (Figs. 4E and F; Supporting

Information Table 1) including (i) four exoglucanases

Figure 2. Activities of extracellular degradative enzymes as

determined by spectrophotometric assays in medium of pure

cultures and co-cultures with litter. (A) Protease activity on

azocasein at pH 7.0. (B) Cellulase activity on carboxymethyl

cellulose at pH 5.0. (C) Pectinase activity on apple pectin at pH

5.0. Controls refer to enzyme activities in sterile medium with

litter; these values were subtracted from activities in culture

supernatants. Data represent means of four independent

experiments. Error bars indicate standard deviation.

Proteomics 2010, 10, 1819–1830 1823


(EC 3.2.1.91; AN0494, AN1273, AN5176, AN5282); (ii) ten

endoglucanases (EC 3.2.1.4, AN1285, AN1602, AN2388,

AN3418, AN3860, AN5320, AN6428, AN8068, AN9524,

AN19419), and (iii) six b-glucosidases (EC 3.2.1.21, AN2217,

AN2227, AN2828, AN4102, AN5976, AN8401). The

production of these enzymes increased over time in pure

cultures of A. nidulans and in co-cultures with P. caroto-vorum. Only four of the A. nidulans cellulases were identified

at low expression levels in the control cultures with glucose

(Fig. 4D), which is in agreement with the zymogram

analyses (Fig. 3).

3.6 Pectinases

Several pectinases were identified when P. carotovorum was

grown on litter as carbon source in the absence and

presence of the fungus (Figs 4H and I; Supporting Infor-

mation Table 1). One pectin lyase (EC 4.2.2.10;

gi|50121327), a not further characterized pectin-degrading

protein (gi|50121327), one exo-poly-a-D-galacturonosidase

(EC 3.2.1.82, gi|50122034), and three different pectate lyases

(EC 4.2.2.2; gi|50122987, gi|50122988, gi|50122989) were

identified. These proteins were expressed at higher levels in

the early growth stage (day 3).

In the experiment with glucose as carbon source, the

pectinase expression pattern was slightly different (Fig 4G).

Three additional bacterial pectinases were detected, two

pectate lyases (EC 4.2.2.2; gi|50120033, 50121477) and one

endo-polygalacturonase (EC 3.2.1.15; gi|50120034).

However, neither the pectin-degrading protein (gi|50121327)

nor the exo-poly-a-D-galacturonosidase (gi|50122034) was

observed.

For pectinases found under both growth conditions, the

peptide counts were generally higher in glucose cultures

(Figs. 4G–I; Supporting Information Table 1). Importantly,

cell numbers were similar in both experiments (data not

shown). Given that the number of peptides belonging to

pectin-degrading enzymes in cultures with litter was

remarkably lower than in glucose cultures, the apparent lack

of pectinolytic enzymes in zymograms of P. carotovorumcultures might result from pectinase expression levels in

litter below the detection limit of the gel-based activity assay.

Growth experiments with different carbon sources indicated

that P. carotovorum grew well on pectin as carbon source

(Fig. 5). This finding might explain why P. carotovorum did

not express cellulolytic enzymes when grown on leaf litter.

A total of ten fungal pectinases were identified in cultures

with beech litter but were absent from cultures in glucose

(Figs. 4G–I). The expression levels increased over time.

There were six pectate lyases (EC 4.2.2.2; AN0741, AN2537,

AN2569, AN6106, AN7646, AN8453), two pectin lyases

(EC 4.2.2.10; AN2331, AN4882), two exo-polygalacturonases

(EC 3.2.1.67, AN8761, AN8891), and one pectin-esterase

(EC 3.1.1.11; AN3390). These findings are in agreement

with the zymogram analysis, which revealed a range of

pectinolytic enzymes in A. nidulans cultures grown on leaf

litter (Fig. 3C).

4 Discussion

Various proteomics analyses of the secretome of plant cell

wall degrading ascomycetes [38], basidiomycetes (e.g. [39, 40]),

and bacteria (e.g. [41]) have been published. However, to the

best of our knowledge, the semi-quantitative proteomics

Figure 3. Zymogram analysis of (A) proteases, (B) cellulases, and (C) pectinases. Cultures were grown in minimal medium containing

either 0.2% glucose or 1% beech litter as sole carbon source. Extracellular enzymes in the medium were analysed after 3 and 17 days.



results presented here are the first on a microbial decom-

position system consisting of more than a single organism.

Our proteomics approach enabled us to identify almost

the entire range of microbial extracellular degradative

enzymes present in cultures with leaf litter, to link

measured enzyme activities to the respective enzymes and

source organisms in mixed culture, and to assess the

enzymatic contributions of a bacterium and fungus to the

degradation of litter constituents.

In accordance with the well-known capability of ascomy-

cetes, including members of the genus Aspergillus to decom-

pose leaf litter [4, 5], our results show that A. nidulans is

capable of degrading a range of plant polymers in leaf litter by

expressing a range of hydrolytic enzymes, including proteases,

cellulases, pectinases, xylanases, mannanases, and rhamno-

galacturonases (Supporting Information Table 1). That

A. nidulans expresses such a wide variety of polymer-degrad-

ing enzymes is not surprising given that efficient hydrolysis of

complex polymers requires synergistic enzymatic interactions

to (i) facilitate access to polymers, such as cellulose, that are

embedded in a heterogeneous matrix [42] and (ii) to cleave

different kinds of linkages present in various polymers such

as cellulose [43, 44], hemicelluloses [45, 46], and pectin

[47, 48]. Our proteomics approach identified ten endocellu-

lases and four exocellulases of fungal origin, which synergis-

tically degrade cellulose, as well as 6 b-glucosidases that

hydrolyse cellobiose, which interferes with cellobiohydrolase

expression (catabolite repression) [49].

Figure 4. Semi-quantitative proteome analysis at day 3 in glucose cultures and day 3 and 17 in litter cultures. Black and white bars

represent peptide counts in pure and co-culture, respectively. (A–C) Proteases. (D–F) Cellulases. (G–I) Pectinases. Data represent the

number of unique peptides assigned to each protein by the Scaffold software; numbers of peptides correlate with protein abundance. For

better comparability, Y-axes of panels representing data from the same enzymes have the same scale. A. nidulans proteins are shown in

the left area of the figures, P. carotovorum proteins are shown in the right area. Codes refer to the accession numbers of the proteins in

the NCBInr database. Details of the proteins and the raw data of peptide count-based quantitation are given in Supporting Information

Tables 1 and 2.

Proteomics 2010, 10, 1819–1830 1825


While fungal enzyme expression and activity were clearly

induced in cultures with beech litter (Figs. 2, 4E, and F;

Supporting Information Table 1), the fungus produced

almost no enzymes when grown on glucose, suggesting that

glucose inhibits expression of polymer-degrading fungal

enzymes (Fig. 4). Our results are in accordance with the

finding that expression of xylanases and cellulases in

Aspergillus sp. depends on the transcriptional activator XlnR

[50, 51], which is activated by low levels of xylose that is

present in decomposing leaf litter [18]. However, high

concentrations of xylose or glucose activate the carbon

catabolite repressor CreA, which, in turn, represses the

expression of a wide variety of hydrolytic enzymes [52, 53].

In contrast to A. nidulans, P. carotovorum only produced

small amounts of pectinases when grown on leaf litter, and

no cellulases or proteases were identified. Pectobacterium is

well known for its arsenal of pectinolytic enzymes, which

are thought to contribute to its pathogenic potential [54]. In

accordance with this information, the present results indi-

cate that P. carotovorum when growing on leaf litter prefer-

entially utilizes pectin, which is a polymer relatively easy to

degrade. Cultivation of P. carotovorum on four different

substrates (Fig. 5) also revealed that the bacterium grows

well on pectin and that it is unable to utilize cellulose as sole

carbon source, although its genome encodes for several

cellulolytic enzymes [16].

Production of extracellular enzymes by Pectobacterium is

influenced by various factors (reviewed in [55]). Many

hydrolytic enzymes (including pectinases) are mainly

induced by (i) polymer degradation products that interfere

with the activity of the transcriptional repressor KdgR

[41, 56], (ii) high cell density (quorum sensing, QS, [57]),

and (iii) anaerobiosis [58] or (iv) low iron concentrations [59].

Expression of hydrolytic enzymes is repressed by (i) cAMP-

controlled catabolite repression in the presence of glucose or

pectin degradation products [60], (ii) nitrogen limitation,

and (iii) elevated temperature [61]. The observed high

expression levels of many extracellular degradative enzymes

in the late stationary phase of P. carotovorum growing on

glucose was probably due to (i) high cell densities inducing

the QS regulatory system(s) and (ii) a lack of catabolite

repression since glucose was entirely consumed at that point

in time. This hypothesis is further supported by the obser-

vation that the highest enzyme expression levels were

observed at day 3 (Fig. 4G, Supporting Information Table 1)

when glucose was detected only in trace concentrations

(data not shown).

Why does P. carotovorum produce only very low amounts

of pectinases and no cellulase and protease in the cultures

with litter as carbon source even though it is able to grow

well on this substrate? That P. carotovorum failed to reach

sufficient cell densities when grown on litter is unlikely

since concentrations of N-acyl homoserine lactones were

high in the culture medium (data not shown). Another

possible explanation is catabolite repression by litter degra-

dation products, which may keep expression levels of

extracellular hydrolytic enzymes low, but still high enough

to support bacterial growth. When the bacteria enter

stationary phase and lower their carbon demand for anabolic

processes, enzyme production might be further repressed.

However, since the interplay of various regulatory systems

involved in cell-wall degrading enzymes is complex, addi-

tional mechanisms might also be effective.

The observed interactions between P. carotovorum and

A. nidulans in co-cultures with litter as the carbon source are

in accordance with several recent studies that have investi-

gated fungal-bacterial interactions during leaf litter decom-

position in aquatic [7, 62–64] and terrestrial [65] model

systems as well as in natural soil habitats [66]. As with our

simple model system, these studies found that fungal

growth was negatively affected by bacteria, whereas the

presence of a fungus promoted bacterial growth. In contrast,

synergistic interaction between fungi and bacteria during

litter degradation have only been reported in very rare cases,

e.g. for fungi and bacteria inhabiting streambed beech litter

microcosms [67]. Encouraged by these consistent findings

we believe that our model system, at least in part, mimics

complex natural communities and will thus help to shed

light on the bacterial and fungal interplay during leaf litter

degradation. Most of these reports could only speculate

about the factors causing the observed antagonistic effects,

such as suppression of fungal growth rather than inter-

ference with fungal enzyme activity [63] or bacterial

competition for carbon and nitrogen [64]. Our proteomics

approach, in contrast, enabled us to pinpoint the molecular

mechanism behind fungal growth inhibition and bacterial

growth stimulation.

Stimulation of bacterial growth by the fungus is most

probably due to the fact that bacteria, apart from actino-

mycetes [68], are unable to physically penetrate the leaf

tissue. Our enzyme assays and semi-quantitative proteome

Figure 5. Growth of P. carotovorum on different carbon sources

in liquid culture. Cell densities were determined by measuring

optical density (OD) of the cultures at 600 nm. Data represent

means of three independent experiments. Error bars indicate

standard deviation.



analyses showed no differences in enzyme activity or

abundance when pure cultures of P. carotovorum grown on

litter were compared with co-cultures where the fungus was

present as well. Clearly, the bacterium did not (or not greatly

in the case of pectinases) contribute to the degradation of

the tested plant polymers. Rather it appeared to profit from

fungal degradation products as resources for growth and

maintenance metabolism. Additionally, litter disintegration

by the fungus probably facilitated access of bacteria to the

inner leaf tissue and hence hydrolysis and use of less

recalcitrant plant polymers such as pectin.

Our results indicate that the reduction of fungal biomass in

co-cultures with P. carotovorum (Fig. 1B) was caused by rapid

bacterial assimilation of litter degradation products released

by fungal enzymatic activity. Since the bacterium can grow

significantly faster than A. nidulans, it should successfully

compete with the fungus for these resources. The observed

decrease in fungal biomass in the presence of P. carotovorum(Fig. 1) did not reduce cellulase and pectinase expression rates

(Fig. 4), indicating that fungal production of degradative

enzymes is increased by nutrient competition. Most interest-

ingly, the presence of the bacterium reduced fungal protease

activity maybe by a reduction of A. nidulans protease expres-

sion (Figs. 2 and 4A). This could imply that P. carotovorum by

some means influences the protease expression by A. nidu-lans, perhaps to protect itself against proteolysis.

Decreased fungal proteolytic activity might lower nitrogen

availability in the litter and thereby limit fungal growth. Our

finding that P. carotovorum grows much better on pectin than

on cellulose as sole carbon source (Fig. 5) together with the

observation that the fungus expresses more protease in co-

culture may suggest that the organisms compete more

strongly for nitrogen (because of reduced fungal protease

activity in presence of the bacterium) than for carbon.

A. nidulans in co-culture with P. carotovorum on litter also

expresses an N,O-diacetylmuramidase (Supporting Informa-

tion Table 1) capable of degrading bacterial cell-walls, which

appears to be a response to the bacterial competitor.

The observation that P. carotovorum grew at the expense

of the fungus during litter decomposition is an example of

‘‘cheating behaviour’’. ‘‘Cheaters’’ are defined as members

of a community that gain advantage of growth in mixed

communities without costs [69]. Allison [70] proposed that

competition for nutrients between effective decomposers

and cheaters is one factor, together with enzyme diffusion

rates and nitrogen limitation, that significantly constrains

enzymatic polymer degradation in natural environments.

Thus, bacterial competition for nutrients and inhibition of

fungal protease activity, as observed in our experiments,

may play an important role in ecosystem functioning.

Our approach involves several drawbacks that limit

straightforward extrapolation of our results to situations in

natural environments. Most important are the low microbial

diversity in our experiments and the growth conditions we

imposed in liquid culture with minimal medium and leaf

litter added. Nevertheless, our results clearly shed new light

on the interactions between fungi and bacteria during litter

decomposition, revealing the key role of a fungus as

decomposer and a bacterium as a cheater that takes advan-

tage of fungal degradative activity, at least under the applied

aerobic culture conditions. However, the scenario might be

completely different when leaf litter has to be degraded

under anaerobic conditions. While aerobic bacteria and

fungi generally produce copious quantities of ‘‘free’’

enzymes, anaerobic bacteria (e.g. Clostridia) but not anae-

robic fungi employ a highly sophisticated cell-bound

enzyme-machinery for the degradation of recalcitrant crys-

talline components of the plant cell wall, which enables

them to out-compete less effective competitors [71, 72];

consequently, the major populations of microorganisms

involved in polymer degradation under anaerobic conditions

are of bacterial origin [73].

Moreover, our results highlight the potential of meta-

proteome analyses to assign degradative enzymes involved

in litter degradation to specific microbial decomposers. This

approach has much to offer for future studies on the

contribution of bacteria and fungi to litter decomposition

under more realistic environmental conditions.

This work is a contribution from the Austrian research networkMICDIF and was supported by the Austrian Science Foundation(FWF). We would like to thank Josef Straus, Institute for AppliedGenetics and Cell Biology, University of Natural Resources,University of Vienna, and Katja Sterflinger, Department ofBiotechnology, University of Natural Resources, University ofVienna, for supplying the A. nidulans strain and Felix Keller,Institute of Plant Biology, University of Zurich for the possibilityto perform carbohydrate analyses. The authors would like to thankAlexander Grunau for critically reading the manuscript.

The authors have declared no conflict of interest.

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Proteome analysis of fungal and bacterial involvement in leaf litter decomposition