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Soil Biology & Biochemistry 39 (2007) 1631–1644

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A microcosm experiment to evaluate the influence of locationand quality of plant residues on residue decompositionand genetic structure of soil microbial communities

Bernard Nicolardota,�, Lamia Bouziria,b,1, Fabiola Bastiana,b, Lionel Ranjardb,��

aINRA—Unite d’agronomie de Laon-Reims-Mons, 2 Esplanade Roland Garros, BP 224, 51686 Reims Cedex 2, FrancebINRA-Universite de Bourgogne, UMR Microbiologie et Geochimie des Sols, CMSE, 17, rue Sully, B.V. 86510, 21065 Dijon Cedex, France

Received 2 November 2006; received in revised form 8 January 2007; accepted 17 January 2007

Available online 21 February 2007

Abstract

The effects of location (soil surface vs. incorporated in soil) and nature of plant residues on degradation processes and indigenous

microbial communities were studied by means of soil microcosms incubation in which the different soil zones influenced by

decomposition i.e. residues, soil adjacent to residues (detritusphere) and distant soil unaffected by decomposition (bulk soil) were

considered. Plant material decomposition, organic carbon assimilation by the soil microbial biomass and soil inorganic N dynamics were

studied with 13C labelled wheat straw and young rye. The genetic structure of the community in each soil zone were compared between

residue locations and type by applying B- and F-ARISA (for bacterial- and fungal-automated ribosomal intergenic spacer analysis)

directly to DNA extracts from these different zones at 50% decomposition of each residue. Both location and biochemical quality

affected residue decomposition in soil: 21% of incorporated 13C wheat straw and 23% left at the soil surface remained undecomposed at

the end of incubation, the corresponding values for 13C rye being 1% and 8%. Residue decomposition induced a gradient of microbial

activity with more labelled C incorporated into the microbial biomass of the detritusphere. The sphere of influence of the decomposing

residues on the dynamics of soluble organic C and inorganic N in the different soil zones showed particular patterns which were

influenced by both residue location and quality. Residue degradation stimulated particular genetic structure of microbial community

with a gradient from residue to bulk soil, and more pronounced spatial heterogeneity for fungal than for bacterial communities. The

initial residue quality strongly affected the resulting spatial heterogeneity of bacteria, with a significance between-zone discrimination for

rye but weak discrimination between the detritusphere and bulk soil, for wheat straw. Comparison of the different detrituspheres and

residue zones (corresponding to different residue type and location), indicated that the genetic structure of the bacterial and fungal

communities were specific to a residue type for detritusphere and to its location for residue, leading to conclude that the detritusphere

and residue corresponded to distinct trophic and functional niches for microorganisms.

r 2007 Elsevier Ltd. All rights reserved.

Keywords: Biodegradation; Crop residues; Detritusphere; Microbial communities; Microcosms; Carbon 13; ARISA

1. Introduction

Soil tillage induces a lot of physical, chemical andbiological changes in soil. Important modifications in soil

e front matter r 2007 Elsevier Ltd. All rights reserved.

ilbio.2007.01.012

ing author. Tel.: +333 26 77 35 83; fax: +33 3 26 77 35 91.

Corresponded to. Tel.: +333 80 69 30 88;

32 24.

esses: [email protected] (B. Nicolardot),

inra.fr (L. Ranjard).

ress: Laboratoire de Traitement et Recyclage des Eaux

m Lif, Tunisia.

processes are due to the location of crop residues in the soilprofile with crop residues remaining at the soil surface inno till and incorporation of residues into the soil byploughing in conventional tillage. In the long term, no-tillsystems generally lead to larger organic C and N stocks inthe soil (Holland and Coleman, 1987; Balota et al., 2004)which mainly accumulate in the top few centimetres(Carter and Rennie, 1982; Salinas-Garcia et al., 2002).Less decomposition occurs in crop residues left at thesoil surface than when soil incorporated (Holland andColeman, 1987; Curtin et al., 1998; Coppens et al., 2006).

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ARTICLE IN PRESSB. Nicolardot et al. / Soil Biology & Biochemistry 39 (2007) 1631–16441632

One main factor might be the climatic conditions in themulch layer where alternate drying and moistening periodsmay restrict microbial activity compared to the perma-nently moist conditions existing around incorporatedresidues (Coppens et al., 2006).

Elsewhere, the number and amounts of microorganisms(i.e. soil microbial biomass) and their distribution withinthe soil profile are modified by tillage practices (Kandeleret al., 1999; Balota et al., 2004). Soil microbial diversity orcomposition may also be changed by no till compared toconventional tillage (Doran, 1980; Lupwayi et al. 2001a;Gomez et al., 2004). Several authors have suggested thatthe residue mulch is mainly decomposed by the fungalcommunity since soil under no tillage contained largeramounts of fungi and larger proportions of fungi tobacteria than under conventional tillage (Holland andColeman, 1987; Frey et al., 1999; Guggenberger et al.,1999; Six et al, 2002). These mulch-associated fungi carryout a simultaneous bidirectional translocation of soil-derived N and mulch-derived C. Most of this mulch-derived C can subsquently be located in the soil (Frey et al.,2003). Bacteria are generally considered to be thepredominant decomposers of incorporated crop residuesunder conventional tillage (Holland and Coleman, 1987;Frey et al., 1999).

Most published work on the effect of location of theorganic matter on its subsequent decomposition hasinvolved large-scale field experiments and either the wholesoil profile or different soil layers (e.g. Biederbeck et al.,1997; Jackson et al., 2003; Gomez et al., 2004; Zhang et al.,2005). Soil heterogeneity has rarely been taken intoaccount except in Lupwayi et al. (2001b). Moreover, theeffect of residue location could not be evaluated because ofthe different and uncontrolled climatic conditions in thevarious experimental tillage systems. Few studies haveinvestigated the effect of residue location on degradationunder controlled conditions (Coppens et al., 2006). Oneinteresting approach was developed by Gaillard et al.(1999) under controlled conditions in which soil hetero-geneity was considered in relation to distance from thepoint source of organic substrate.

Many of the works focussed on dynamics and structureof the soil microbial community in tillage systems onlyconsidered the bacterial and/or fungal biomass (e.g. Entryet al., 2003; Lupwayi et al., 1999; Ibekwe et al., 2002; Fenget al., 2003; Spedding et al., 2004) and physiologicaldiversity (e.g. Gomez et al., 2004; Lupwayi et al., 1998,2001b; Zhang et al., 2005). As a result, most such studiestended to consider the entire microbial community as afunctional ‘‘black box’’ whereas microbial communities arecomplex assemblages of species, with different metaboliccharacteristics and physiological requirements, and eachone driving at least one of the multiple reactions of organicmatter transformation. In this context, a major challenge inmicrobial ecology has been to precisely identify thosemicrobial populations and activities involved in carbondegradation and the reciprocal structuration of soil

communities according to carbon source. New molecularmethods based on the analysis of nucleic acids directlyextracted from the soil matrix have been developed overthe past ten years. These circumvent the limitationsresulting from the selectivity and unrepresentativity ofculture-based methods and have revealed new aspects ofsoil microbial diversity (for review see Ranjard et al., 2000).However, despite these methodological developments, fewstudies have looked at the genetic structural dynamics ofthe soil microbial community during plant residue decom-position (Lundquist et al, 1999) and under the effect ofmulching (Tiquia et al., 2002).Thus, our main objectives were to study the effects of the

location of various types of plant residues on the structureof the soil microbial community involved in their decom-position. This would consist of highlighting the significantdifferences in soil microbial community structure whichwere induced by the initial quality of the carbonaceoussubstrate and its initial location in the soil. Our study wasperformed under controlled conditions and considered thedifferent soil zones (residues, detritusphere and bulk soil)that were variously influenced by the decompositionprocesses. 13C-labelled plant materials were used to studyresidue decomposition and the assimilation of organiccarbon by the soil microbial biomass. Microbial biomass ineach location was measured by fumigation extractionmethod. This has been widely used as a bioindicator toevaluate the effect of different agricultural practices on soilmicroorganisms (Marschner et al., 2003). The geneticstructures of the bacterial and fungal communitiesindigenous to the different soil and residue zones underthe different incubation conditions were assessed by theDNA ‘‘fingerprinting approach’’, known as automated-ribosomal intergenic spacer analysis (ARISA).

2. Materials and methods

2.1. Soil type and preparation

The soil was an Orthic Luvisol collected at the Estrees-Mons INRA experimental station (49.531N, 3.011E) andsampled from the 0–15 cm layer. Calibrated soil aggregates(2mm diameter) were obtained by sieving the soil sampleand were stored at 4 1C until use. The main soilcharacteristics are presented in Table 1.

2.2. Plant residues

Two different plant residues were used in this study:mature wheat straw (Triticum aestivum) and young ryeleaves (Secale cereale L.). Both were labelled with 13C bygrowing the plants in a labelling chamber. The growthconditions are detailed in Aita et al. (1997) and Coppens(2005), respectively. The plants were dried at 80 1C and cutinto pieces 1 cm in length. The C and N contents and 13Cisotopic abundance were determined with an NA 1500elemental analyser (Fisons, Milan, Italy) coupled with an

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Table 1

Main characteristics of soil used for incubation

Characteristics Unit Value

Clay (o2mm) g kg�1 185

Silt (2–20mm) g kg�1 296

Silt (20–50mm) g kg�1 457

Sand (50–200mm) g kg�1 57

Sand (200mm–2mm) g kg�1 5

Organic C g kg�1 8.31

Total N g kg�1 0.99

C:N ratio 8.4

CaCO3 g kg�1 3.1

pH (H2O) 8.0

Microbial biomass C mgkg�1 178

Table 2

Analytical characteristics of plant materials

Characteristics Unit Rye plants Wheat straw

C g 100 g�1 d.m. 44.20 44.47

N g 100 g�1 d.m. 2.14 0.67

Isotopic excess Atoms 13C 100 atoms�1 2.88 1.99

Water-soluble C g 100 g�1 C 37.5 11.1

Van Soest fractions

Soluble g 100 g�1 d.m. 52.7 23.3

Hemicellulose g 100 g�1 d.m. 24.7 28.7

Cellulose g 100 g�1 d.m. 21.2 42.8

Lignin g 100 g�1 d.m. 1.4 5.2

B. Nicolardot et al. / Soil Biology & Biochemistry 39 (2007) 1631–1644 1633

Isochrom mass spectrometer (Fisons, Manchester, UK).Water-soluble C was obtained by extracting 50mg ofdried plant material in 25mL water (20 1C, 30min), andanalysing the dissolved C with a 1010 TOC analyser (OIAnalytical, College Station, TX). The soluble, hemicellu-lose, cellulose and lignin fractions were determined usingthe methods described by Lineres and Djakovitch (1993)and Van Soest (1963). The analytical characteristics of theplant materials are presented in Table 2.

2.3. Treatments, preparation of soil cores and incubation

Decomposition of the plant materials was studied inrepacked soil cores (7 cm diameter, 4.8 cm height) obtainedby uniaxial confined compression of a mass of calibratedwet aggregates in a cylindrical mould (Fazzolari et al.,1998) (5 replicates for each experimental treatment andsampling date). For residues left at the soil surface, theequivalent of 268.4 g dry calibrated soil aggregates was firstintroduced into the mould and compacted to obtain a finalbulk density of 1.45. The equivalent of 1.0 g dry plantresidues was then introduced into the mould and dis-tributed on the core surface. The soil core and residueswere then compacted together to give a final bulk densityof 1.45. For the soil-incorporated residues, half the mass ofcalibrated soil aggregates (equivalent to 134.2 g dry soil)was first introduced in the mould and compacted to obtain

a final bulk density of 1.45. The equivalent of 1.0 g dryplant residues was then introduced into the mould anddistributed on the surface of this half core, after which therest of the calibrated soil aggregates (equivalent to 134.2 gdry soil) was introduced into the mould to cover the plantresidues. The soil aggregates and residues were thencompacted together to obtain a final bulk density of 1.45.Soil cores with no addition of plant residues were alsoconsidered (control soil, 5 replicates for each experimentaltreatment and sampling date).The soil moisture content was fixed and maintained at a

matrix potential of �0.05MPa by adding deionised water.No inorganic N was added to the soil before incubation.Each soil core was placed in a 3L airtight glass jar with aCO2 trap (30mL 1M NaOH). The soil cores wereincubated at 1570.3 1C for 168 days. The jars wereopened periodically to renew the atmosphere and replacethe CO2 traps.

2.4. Soil sampling and determinations

Three zones were considered for each sampling date:plant residues, detritusphere (6mm thick soil layer adjacentto the plant residues, as suggested in Gaillard et al., 2003)and bulk soil (soil below the 6mm detritusphere). First, theremaining plant residues were carefully separated from thesoil with tweezers after opening the core along the strawlayer in the case of incorporated plant residues. Then, thedetritusphere and bulk soil were separated by sectioningthe soil cores and half cores (incorporated residuetreatments) with a cutting device (Millon, 2004). In theincorporated residues treatments, both detritusphere layerswere put together as well as both bulk soil layers. Each soilzone (bulk soil and detritusphere) were then homogeneizedand sampled for further determinations.The dry mass of the remaining plant residues (2 replicates/

date) was determined after drying for 24h at 80 1C. The Ccontent and 13C isotopic abundances of the remaining plantresidues, detritusphere and bulk soil (2 replicates/date) weredetermined using an NA 1500 elemental analyser coupledwith an Isochrom mass spectrometer. Water-soluble C wasextracted from the detritusphere and bulk soil (dry soil:waterratio ¼ 1/5; agitation 30min at 20 1C, 2 replicates/date) anddetermined using the 1010 TOC analyser. The 13C isotopicabundances of the freeze-dried extracts were then determinedusing the elemental analyser coupled with the mass spectro-meter. Microbial biomass C and 13C in the detritusphere andbulk soil were determined after measuring soluble organic Cin 0.025M K2SO4 extracts of CHCl3 fumigated and non-fumigated soil (dry soil:extractant ratio ¼ 1/5; agitation30min at 20 1C, 3 replicates/date) (Trinsoutrot et al.,2000a). Inorganic N was extracted from the detritusphereand bulk soil with 1M KCl (dry soil:KCl ratio ¼ 1/5–3/10;30min at 20 1C, 2 replicates/date), and NHþ4 �N and NO�3 �Nwere determined by continuous flow colorimetry with anauto-analyser (TRAACS 2000, Bran & Luebbe, Norderstedt,Germany) using adaptations of the methods proposed by


Kamphake et al. (1967) and Krom (1980). The CO2 and13CO2 collected in NaOH traps (5 replicates) were deter-mined using the techniques described in Trinsoutrot et al.(2000a). Finally the plant residues, detritusphere and bulksoil samples (3 replicates/date) used for DNA extraction werestored at �40 1C prior to analysis.

2.5. Extraction and purification of total DNA from soil

zones and plant residues

Microbial DNA was extracted from independent tripli-cates of the plant residue, detritusphere and bulk soil at the

0 30 60

mg

13 C

-resid

ue k

g-1

soil

0

300

600

900

1200

1500

Fig. 1. Amounts of 13C labelled plant residues remaining in soil for the diff

0 30 60

mg 1

3C

-CO

2 k

g-1

soil

0

200

400

600

800

1000

1200

W

W

R

R

Fig. 2. Cumulative evolution of 13C labelled CO2 for the different e

sampling time corresponding to degradation of half theresidues (50% dry mass loss during the 168-day incuba-tion ¼ about 15 days for rye leaves and 1 month for wheatstraw) according to the method described by Ranjard et al.(2003). Briefly, 1 g from each soil and residue sample wasmixed with 4mL of a solution containing 100mM Tris(pH 8.0), 100mM EDTA (pH 8.0) 100mM NaCl, and 2%(wt/vol) sodium dodecyl sulfate. Two grammes of 106 mm-diameter glass beads and 8 glass beads of 2-mm diameter,were added to the mixture in a bead-beater-tube.The samples were then homogenized for 30 s at 1600 rpmin a mini bead-beater cell disruptor (Mikro-dismembrator

days

90 120 150 180

Wheat straw incorporated

Wheat straw surface

Rye leaves incorporated

Rye leaves surface

erent experimental treatments. Bars indicate standard deviation values.

days

90 120 150 180

heat straw incorporated

heat straw surface

ye leaves incorporated

ye leaves surface

xperimental treatments. Bars indicate standard deviation values.

ARTICLE IN PRESSB. Nicolardot et al. / Soil Biology & Biochemistry 39 (2007) 1631–1644 1635

S. B. Braun Biotech International). The samples wereincubated for 20min at 70 1C, then centrifuged at 14 000g

for 1min at 4 1C. The collected supernatants wereincubated for 10min on ice with 1/10 volume of 3Mpotassium acetate (pH 5.5) and centrifuged at 14 000g for5min. After precipitation with one volume of ice-coldisopropanol, the nucleic acids were washed with 70%ethanol. DNA was separated from the residual impurities,particularly humic substances, by centrifuging through twotypes of minicolumns. Aliquots (100 mL) of crude DNAextract were loaded onto PVPP (polyvinyl polypyrroly-done) minicolumns (BIORAD, Marne-la-Coquette,France) and centrifuged at 1000g for 2min at 10 1C. Thecollected eluate was then purified with the Geneclean turbokit (Q-Biogene, Illkirch, France).

2.6. Automated-RISA fingerprinting

The genetic structure of the microbial communities wasdetermined by ARISA, which exploits the variability inlength of the intergenic spacer (IGS) between small (16S forbacteria and 18S for fungi) and large (23S for bacteria and28S for fungi) subunit rRNA genes in the rrn operon. Thebacterial and fungal ribosomal IGS were amplified withthe primers S-D-Bact-1522-b-S-20/L-D-Bact-132-a-A-18



days

mg 1

3C

-solu

ble

kg

-1 s

oil

0

1

2

3

4

5

mg 1

3C

-solu

ble

kg

-1 s

oil

0

1

2

3

4

5

14 28 56 84 168

Fig. 3. Distribution of 13C labelled water soluble C in the different soil zones

whole soil taking into account the proportion of soil zones (detritusphere repres

incorporated residues). Bars indicate standard deviation values.

and ITS1F/3126T, respectively, with PCR conditions asdescribed by Ranjard et al. (2003). Fifty nanogrammes ofDNA were used as template in PCR. A fluorescent-labelledprimer was used for the LiCors DNA sequencer (Scien-ceTec, Les Ulis, France) in B-ARISA and F-ARISA,namely the IRD 800 dye fluorochrome (MWG SA Biotech,Ebersberg, Deutschland). PCRs were performed usingthe S-D-Bact-1522-b-S-20 and 3126T primers labelled attheir 50 end with the IRD800 fluorochrome. The concen-tration of labelled PCR products was estimated, andbetween 0.5 and 1 mL of the product was added todeionised formamide and denatured at 90 1C for 2min.ARISA fragments were resolved on 3.7% polyacrylamidegels and run under denaturing conditions for 15 h at3000V/60W on a LiCors DNA sequencer (ScienceTec).The data were analysed using the 1D-Scan software(ScienceTec). This software converted fluorescence datainto electrophoregrams in which the peaks representedthe PCR fragments. The heights of the peaks werecalculated together with the median filter option andthe Gaussian integration in 1D-Scan, and representedthe relative proportion of fragments in the total pro-ducts. Lengths (in base pairs) were calculated byusing a standard size with bands ranging from 200 to1206 bp.

Wheat straw surface

mg 1

3C

-solu

ble

kg

-1 s

oil

Detritusphere

Bulk soil

Rye leaves surface

days

14 28 56 84 168

0

1

2

3

4

5

mg 1

3C

-solu

ble

kg

-1 s

oil

0

1

2

3

4

5

for the different experimental treatments. Values are calculated for 1 kg of

ents, respectively, 12.5% and 25% of soil weight for residues at surface and


2.7. Statistical analysis

The statistical analyses of the C and N data wereperformed using the ANOVA procedure of SAS software(SAS Institute, 2001), with a means comparison byStudent–Newman–Keuls test (pp0.05). Significant differ-ences (pp0.05) in DNA yields between the different samplesizes in each soil were determined using Statview-SEsoftware and Student’s t-test.

Data obtained from the 1D-Scan software were con-verted into a table summarizing the band presence (i.e.


mg inorg

anic

-N k

g-1

soil

Detritusphere

Bulk soil


mg inorg

anic

-N k

g-1

soil

0 14 28 56 84 168

Control soil

days

mg inorg

anic

-N k

g-1

soil

0

1

2

3

4

5

6

0

10

20

30

40

50

60

0

10

20

30

40

50

60

Fig. 4. Distribution of inorganic N in the different soil zones for the treatme

control soil. Values are calculated for 1 kg of whole soil taking into account the

25% of soil weight for residues at surface and incorporated residues). Bars in

peak) and intensity (i.e. Gaussian area of peak) usingthe PrepRISA programme (Ranjard et al., 2003). A robustanalysis was ensured by integrating 100 bands for B- andF-ARISA profiles with a 2 bp resolution (Ranjard et al.,2003). PCA was carried out using a B-ARISA andF-ARISA covariance matrix on the data matrix (commu-nities as rows and bands as columns). This method enabledthe bacterial or fungal communities to be ordinated byplotting the scores for the first two principal components intwo dimensions. PCA were performed using the ADE-4software (Thioulouse et al., 1997). This ADE-4 software

mg inorg

anic

-N k

g-1

soil

Wheat straw surface

Rye leaves surface

days

0 14 28 56 84 168

mg inorg

anic

-N k

g-1

soil

0

10

20

30

40

50

60

0

1

2

3

4

5

6

nts with input of residues and evolution of inorganic N in whole soil for

proportion of soil zones (detritusphere represents, respectively, 12.5% and

dicate standard deviation values.


depicted the statistical area representing 90% confidencefor the sample replicates on the factorial map.

3. Results and discussion

3.1. Influence of residue quality and location on 13C fluxes

in soil

As expected, rye leaves decomposed more quickly andto a greater extent than wheat straw (Figs. 1 and 2).Differences in decomposition were related to residuequality (Table 2) as reported by several authors (Trinsou-trot et al.; 2000b; Gaillard et al., 2003; Abiven et al., 2005)who showed that decomposition under controlled condi-tions was related to the initial water soluble or solublecompounds present in the plant materials. Wheat strawdecomposition was poorly affected by location, differencesin decomposition being statistically significant for CO2

emission only during the first 14 days of incubation. Incontrast, location did influence the decomposition of ryeleaves and a little more CO2 was emitted when the residueswere superficial (Fig. 2), but more undecomposed ryeleaves remained (po0.01) when the residues were left onthe soil surface (Fig. 1). This apparent discrepency betweenCO2 emission and amounts of remaining residues is



days

mg 1

3C

-bio

mass k

g-1

soil

0

20

40

60

80

100

120

mg 1

3C

-bio

mass k

g-1

soil

0

20

40

60

80

100

120

14 28 56 84 168

Fig. 5. Distribution of 13C labelled soil microbial biomass in the different soil

1 kg of whole soil taking into account the soil proportion of soil zones (detritusp

surface and incorporated residues). Bars indicate standard deviation values.

explained by the fact that CO2 emission corresponds toapparent C mineralization and does take into account theincorporation in microbial biomass (see below). Mostauthors (Douglas et al., 1980; Holland and Coleman,1987; Curtin et al., 1998) have shown that decompositionunder field conditions is enhanced when the residues areincorporated. Similar results were found by Coppens et al.(2006) for laboratory experiments with controlled rainevents. In contrast, when studies involved soil incubationunder controlled conditions, differences between thedecomposition of plant materials at the soil surface or soilincorporated were shown to be weak or null (Cogle et al.,1989; Aulakh et al., 1991). In fact, differences in decom-position between residues incorporated in soil or left at thesoil surface may relate to moisture conditions in the mulchlayer (Aulakh et al., 1991; Coppens et al., 2006), whilemoisture conditions in our study were optimal for residuedecomposition in all experimental treatments. Anotherfactor of influence is the inorganic N availability which willbe discussed later.During biodegradation, water soluble C was transferred

from the decomposing residues to the detritusphere andbulk soil as observed by Frey et al. (2003). Indeed, somewater soluble 13C was continuously present in both soilzones during the incubation period (Fig. 3), but was always

Wheat straw surface

Detritusphere

Bulk soil

Rye leaves surface

days

14 28 56 84 168

mg 1

3C

-bio

mass k

g-1

soil

0

20

40

60

80

100

120

mg 1

3C

-bio

mass k

g-1

soil

0

20

40

60

80

100

120

zones for the different experimental treatments. Values are calculated for

here represents, respectively, 12.5% and 25% of soil weight for residues at


less than 0.1% of the added 13C. The effect of residuelocation on the amounts present in the soil fractions wasstatistically non significant, although a little more labelledsoluble carbon was found in the detritusphere when theresidues were incorporated. Nevertheless, the amounts oflabelled soluble C were significantly higher in the detritu-sphere than in bulk soil and higher for rye leaves. Inaddition the total residue-derived 13C present in thedetritusphere and bulk soil represented to 5.7–19.1% and1.2–2.9% of the initial 13C input at the end of incubation(results not shown). This confirms the spatial limitation ofthe sphere of influence of residues as reported by Gaillard

810 bp

B-ARISA

540 bp

230 bp

325 bp

1051 bp

1532 bp

200 bp

325 bp

540 bp

810 bp

1051 bp

1478 bp

Wheat incorporated Rye surface

R Dsph BS R Dsph BS

105

153

81

54

32

20

Fig. 6. Example of B-ARISA and F-ARISA profiles obtained from DNA extr

bulk soil (BS) in microcosms incubated with wheat straw and rye leaves at soil

by residue degradation.

et al. (1999) who showed that decomposing plant residuesproduced a steep gradient in microbial activity and residue-derived 13C in the soil mainly in the 4-mm zone in contactwith the residue.

3.2. Influence of residue quality and location on soil

inorganic N dynamics in soil

Very low amounts of inorganic N were observed in thesoil zones for wheat straw at both locations (Fig. 4). Wheatstraw incorporation in fact induced net N immobilizationin both zones, although small amounts of N were

F-ARISA

Wheat surface Rye incorporated

R Dsph BS R Dsph BS

1051 bp1 bp

1532 bp2 bp

810 bp0 bp

540 bp0 bp

325 bp5 bp

200 bp0 bp

acted from independent replicates of residue (R), detritusphere (Dsph) and

surface or incorporated. Arrows indicate particular populations stimulated


mineralized at the end of the incubation period. Surface-applied wheat straw induced net N immobilization in thedetritusphere whereas N mineralization was observed forthe bulk soil during the first 14 days followed byprogressive N immobilization from day 14 to day 84.These data confirm the findings of several authors (Aulakhet al., 1991; Coppens et al., 2006) who reported greater Nimmobilization when residues were incorporated. Theinorganic N for rye leaves and both locations, increasedin both soil zones during the incubation period whichindicated net N mineralization, with more N mineralizedthan in the control soil. These dynamics shows that theorganic N in rye leaves is mineralized which is inaccordance with the findings of many other authors(e.g. Trinsoutrot et al., 2000a; Mendham et al., 2004;Abiven et al., 2005) who showed that plant materialswith a low C:N ratio induced net N mineralization duringtheir decomposition in soil. Our results suggest that wheatstraw decomposition was probably limited by the avail-ability of soil inorganic N, as shown by Recous et al.(1995), which could explain the weak effect of location ondecomposition. Finally, comparative N dynamics in thedetritusphere and bulk soil suggest a transfer of inorganicN from bulk soil to detritusphere and vice versa. This mayoccur by physical processes (e.g. diffusion) and biologicaltransfer.

PC2 15.2%

PC1 48.7%

Dsph R

BS

Soil surface

Wheat s

Rye lea

Soil surface

PC2 16.3%

BS RDsphPC1 39.2%

Fig. 7. Principal component (PC1�PC2) plots generated from B-ARISA pro

(BS) in microcosms incubated with wheat straw or rye leaves at soil surface o

3.3. Influence of residue quality and location on 13C

incorporation in the microbial biomass

Significantly more labelled C was present in themicrobial biomass of the detritusphere for both residuesthan in bulk soil (Fig. 5), confirming the conclusion ofGaillard et al. (1999) that microbial assimilation occursessentially in the detritusphere. In fact, the soluble Ctransferred into the soil from decomposing residues israpidly assimilated by soil microorganisms. Thus, thefurther the soil is from the residues, the less C is availableto the soil microorganisms, thereby leading to considerablesoil heterogeneity.More 13C was present in the microbial biomass during

the decomposition of rye leaves than of wheat straw andmore 13C was incorporated into the microbial biomasswhen rye leaves were incorporated (Fig. 5). These resultsindicate a higher rate of decomposition when this residue isincorporated (Fig. 1) and coincide with a higher avail-ability of the residue-derived water-soluble C (Fig. 3). Aneffect of location of the wheat straw residue was onlyobserved in the detritusphere. These results confirm thatrye leaves provide a much more decomposable substratefor soil microorganisms than wheat straw, as observed byTrinsoutrot et al. (2000a) who found that more C wasassimilated by the soil microbial biomass from residues

PC1 64.3%

PC2 18.1%

RDsph

BS

Incorporated

traw

ves

Incorporated

PC1 66.4%

PC2 9.8%

R

Dsph

BS

files obtained from the residue (R), the detritusphere (Dsph) and bulk soil

r incorporated. Ellipses represent 90% confidence limits.


with high N and soluble C contents. The lower incorpora-tion into the microbial biomass of labelled C fromwheat straw is coherent with its lower decomposition(Figs. 1 and 2) and lower availability of released water-soluble C (Fig. 3). The poor effect of wheat residue locationon microbial assimilation confirmed the limited biodegra-dation in both locations, possibly due to soil inorganic Navailability (Fig. 4).

3.4. Influence of residue quality and location on spatial

distribution of the microbial community structure

In this study, the genetic structures of the bacterial(B-ARISA) and fungal (F-ARISA) communities in plantresidues, detritusphere and bulk soil, were compared at asingle stage in degradation during the 168-day incubation,corresponding to a loss of 50% dry mass (about 15 days forrye leaves and 1 month for wheat straw). ARISAfingerprinting of the community, by direct determinationof DNA extracts from the different soil zones, providedcomplex profiles with peaks ranging from 250 bp (i.e.100 bp-IGS) to more than 1000 bp (1050-bp IGS) forbacteria and from 350 to 1000 bp for fungi (Fig. 6). Visualcomparison of the B- and F-ARISA profiles showed thateach soil zone was characterized by a specific pattern,suggesting a particular genetic structure of the community.

PC1 33.3%

PC1 53.8%

PC2 19.8%

R

BS

R

PC2 19.4%

Soil surface

Wheat s

Dsph

BS

Rye lea

Soil surface

Dsph

Fig. 8. Principal component (PC1�PC2) plots generated from F-ARISA pro

(BS) in microcosms incubated with wheat straw or rye leaves at soil surface o

The PCA of B-ARISA confirmed and expressed thesedifferences as a gradient from residue to bulk soil (Fig. 7)suggesting that residue degradation stimulated certainminor populations in the bulk soil and detritusphere (asindicated by arrows in Fig. 6). This spatial gradient invariability confirmed that a decomposing residue results insoil heterogeneity but that its sphere of influence is spatiallylimited (Gaillard et al., 1999). In addition, this spatialheterogeneity exhibits similarities with that observed in therhizosphere between root tissue and bulk soil (Mougelet al., 2006). Altogether, these results confirm that both therhizosphere of living plants and residues of dead plantsrepresent ‘‘hot spots’’ of a readily available energy source,C and nutrients for soil microorganisms, and stronglymodify the genetic structure of the community bystimulating particular populations, especially as the soilsystem is often substrate limited as regards microbialgrowth (Mougel et al., 2006; Lejon et al., 2007).In the case of wheat residue decomposition, the weak

discrimination between indigenous bacterial communitiesin the detritusphere and bulk soil, whatever the residuelocation (Fig. 7), did not coincide with the above-describedspatialisation of the microbial biomass (Figs. 3–5), or withthe previously reported assimilation of decomposingsubstrates (Gaillard et al., 1999; Ronn et al., 1996). Theseauthors recorded spatial heterogeneity in microbial activity

PC1 36%

PC2 20.3%

R

PC1 39.5%

PC2 19.6%

R

BS

Incorporated

traw

Dsph

BS

ves

Incorporated

Dsph

files obtained from the residue (R), the detritusphere (Dsph) and bulk soil

r incorporated. Ellipses represent 90% confidence limits.


around decomposing wheat straw over a millimetre scale,with the area of influence limited to 4mm from theresidue. Altogether, these results suggest a weak relation-ship between the rate of residue decomposition and thecomposition of the community (Lejon et al., 2007).In contrast, a significant discrimination between thebacterial communities in each zone was apparent forrye leaves, and even more so when the residue wasincorporated (Fig. 7). The observed differences betweenrye leaves and wheat straw suggested a strong interactionbetween the initial quality of the residue and resultingspatial heterogeneity, and confirmed the hypothesis ofGaillard et al. (2003). These authors concluded that thedecomposition of young rye leaves led to a higher spatialheterogeneity of the decomposing microorganisms due to agreater release of soluble organic compounds.

A more significant discrimination between bulk soil anddetritusphere was recorded for the fungal community thanfor bacteria (Fig. 8), suggesting that plant residue degrada-tion, whatever the type of residue and location, led to a more

B-ARISA

WIWSRS RI WIWSRS RI

230 bp

325 bp

540 bp

810 bp

1051 bp

8

1

1532 bp 1

Residues Detritusphere

Fig. 9. B- and F-ARISA profiles obtained from DNA extracted from independ

wheat straw at soil surface (WS) or incorporated (WI) and rye at soil surface

significant spatial heterogeneity of fungal genetic structureand thus a more significant impact of the residue in thedetritusphere soil. This might be explained by the shape offungal bodies and the growth patterns produced by thefilamentous ramified mycelia that enable residue-colonizingfungi to radiate through the soil over several millimeters andthus modify the population structure in the detritusphere (deBoer et al., 2005). This hypothesis was supported by visualobservation of the profiles which revealed bands common toresidue and detritusphere but not detected in bulk soil(indicated by arrows in Fig. 6). However, several bands werealso specific to the detritusphere suggesting that particularpopulations were stimulated in this zone possibly by theamount and/or specific quality of soluble carbon (Lejonet al., 2005, 2007). It should be noted that, in our case,residue location had no significant influence on spatialheterogeneity of the fungal community and did not confirmthe possibly less important involvement of fungi in thedecomposition process when residues are incorporated(Holland and Coleman, 1987; Frey et al., 1999).

F-ARISA

WIWSRS RI WIWSRS RI

230 bp

325 bp

540 bp

10 bp

051 bp

532 bp

DetritusphereResidues

ent replicates of residues and detritusphere in microcosms incubated with

(RS) or incorporated (RI).


3.5. Influence of plant residue quality and location on the

microbial genetic structure in each soil zone

The data obtained from the ARISA profiles werecomputed to compare the genetic structures of thecommunities in each zone according to residue type (ryeleaves vs. wheat straw) and location (incorporated vs.surface) (Fig. 9). Whatever the location, PCA revealed asignificant discrimination between the detritusphere of ryeleaves and wheat straw, suggesting that the structures ofthe indigenous bacterial and fungal communities in thedetritusphere were more specific to the type of residue thanto its location (Fig. 10). In soil, microorganisms are mainlyheterotrophic and carbon limited and such discriminationcould result from the contrasting availability and quality ofthe carbon source supplied by wheat straw and rye leavesdecomposition as previously discussed (Fig. 3 and Table 2).A similar hypothesis was tested in forest and agriculturalfield studies where the distinct composition of microbialcommunities was believed to be due to differences in thesoil organic management (Myers et al., 2001; Lejon et al.,2005; Yang et al., 2003) leading to different types ofdissolved organic C in the soil (Marschner et al., 2003).From a functional point of view, discrimination betweenthe community structures can be related to differences in

PC1 44.2%

PC2 22.9%

Residues

PC1 26.1%

PC2 21.6

RIWI

WS

B-ARIS

RS

F-ARIS

RS

RI

WS

WI

Residues

Fig. 10. Principal component (PC1�PC2) plots generated from B- and F-ARI

incubated with wheat straw at soil surface (WS) or incorporated (WI) and rye a

limits.

the rate of decomposition (Figs. 1 and 2), these being moresignificant for residue type than for their location.In the residue zone, PCA revealed a more significant

discrimination of the bacterial and fungal communitiesbetween residue location than between residue type(Fig. 10). Significant discrimination was only observedbetween wheat straw and rye leaves for bacterial commu-nities when the residues were on the surface. Theseunexpected results suggest that similar bacterial and fungalpopulations colonized both residue types and that thephysical location of the residue had a greater influence thanits quality on the microbial composition of colonizers.These data could be partly explained by the stressconditions, in terms of moisture content, generallyobserved in the mulch layer and by the colonizationsurface offered to microorganisms which is reduced whenthe residue is incorporated (Holland and Coleman, 1987;Aulakh et al., 1991; Coppens et al., 2006).

4. Conclusions

Altogether, our results emphasized that the presence ofdecomposing residues in soil induced a gradient from bulksoil to residue site of microbial density, processes and alsoin the composition of the community. In addition, this

PC1 30.4%

PC2 21%

PC2 18.3%

DetritusphereA

RIWS

WIRSPC1 46.1%

A

RS

RI

WSWI

Detritusphere

SA profiles obtained from the residues and the detritusphere in microcosms

t soil surface (RS) or incorporated (RI). Ellipses represent 90% confidence


sphere of influence may be modified by type of residue aswell as its location in the soil. This study represents aninitial step in the ecological study of crop residuedecomposition in soil. Nevertheless, it remained difficultto extrapolate our results in the field conditions consideringagricultural practices such as organic management andtillage. Future research will need to confirm, at the fieldlevel, the influence of residue location on microbialcommunity by taking into account agronomic resultantsuch as soil fertility, carbon turn over as well as ecologicalparameters such as microbial diversity and stability.

Acknowledgements

L. Bouziri and F Bastian received grants, respectively,from INRA and La Region Champagne-Ardenne and theirwork was supported by INRA. We would like to thank G.Alavoine, O. Delfosse, F. Millon, S. Millon and M.J. Herrefor their technical assistance. Thanks are also extended toV. Nowak for her technical help during this study.

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