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Soil Biology & Biochemistry 79 (2014) 132e139

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Soil Biology & Biochemistry

journal homepage: www.elsevier .com/locate/soi lbio

Assimilation and accumulation of C by fungi and bacteria attached tosoil density fractions

Pierre-Joseph Hatton a, *, 1, Samuel Bod�e b, 1, Nicolas Angeli c, Pascal Boeckx b,Bernd Zeller a, S�everine Boiry d, Louisette Gelhaye a, Delphine Derrien a

a INRA-Nancy, Biog�eochimie des �Ecosyst�emes Forestiers, 54280 Champenoux, Franceb Isotope Bioscience Laboratory e ISOFYS, Faculty of Bioscience Engineering, Ghent University, 9000 Ghent, Belgiumc INRA-Nancy, Ecologie et Ecophysiologie Foresti�eres, 54280 Champenoux, Franced IBEB/SBVME Groupe de recherche Appliqu�ee en Phytotechnologie, UMR 7265 CNRS/CEA/Univ. Aix Marseille, CEA de Cadarache, 13108 Saint Paul lezDurance, Cedex, France

a r t i c l e i n f o

Article history:Received 30 April 2014Received in revised form8 September 2014Accepted 13 September 2014Available online 24 September 2014

Keywords:SoilAmino sugars13CDensityLC-IRMSForest

* Corresponding author. Current address: DepartmeBiology, The University of Michigan, Ann Arbor, MI, U

E-mail addresses: pierre-joseph.hatton@nancy.ihatton_pj@yahoo.fr (P.-J. Hatton).

1 Equal contribution of both authors.

http://dx.doi.org/10.1016/j.soilbio.2014.09.0130038-0717/© 2014 Elsevier Ltd. All rights reserved.

a b s t r a c t

Soil microorganisms play a key role in soil organic matter (SOM) dynamics, but little is known about thecontrols affecting the distribution of microbial biomass and their residues in soil. Here, a forestedCambisol topsoil was incubated with 13C-labeled glycine or beech leaves for 12 weeks prior to sequentialdensity fractionation. The incorporation of the 13C label in amino sugars (AS) was used to gain insightinto bacterial and fungal assimilation of the substrates. AS derived from glycine or leaves were comparedto total AS to investigate how microbial residues and active communities were distributed among soildensity fractions.

Bacteria slightly dominated leaf C assimilation, while a pronounced fungal dominance was observedfor glycine. The glycine-derived AS and original AS were similarly distributed among the soil densityfractions, both peaking in microbial aggregates (1.8e2.4 g cm�3). Leaf-derived AS were mostly found inassociation with the plant debris (<1.65 g cm�3). The ratios of substrate-derived AS C to substrate-derived C increased with soil fraction density for both glycine and leaves. The same pattern wasobserved with original AS C to soil fraction C ratios. We concluded that bacteria and fungi were mostactive where the resource was even though their residues accumulate mostly in microbial aggregates (1.8e2.4 g cm�3). We suggest that such accumulation might be attributed to (1) an increasing stabilizationefficiency of microbial residues and (2) the progressive SOM transfer, from plant debris to microbialaggregates (1.8e2.4 g cm�3).

© 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Soil microorganisms are widely recognized as important con-trolling factors for soil organic matter (SOM) dynamics (Schmidtet al., 2011; Zhou et al., 2012), primarily because of their promi-nent role in SOM degradation and accumulation (Kindler et al.,2009; Miltner et al., 2012). Microbial residues constitute animportant component of SOM (Koegel-Knabner, 2002; von Lützowet al., 2006; Koegel-Knabner et al., 2008) and their allocation might

nt of Ecology & EvolutionarySA.nra.fr, pjhatton@umich.edu,

well play a key role long-term soil C sequestration (Schimel andSchaeffer, 2012). By jointly analyzing fumigation-extracts and bio-markers of total and living microbial biomasses, Appuhn andJoergensen (2006) showed that half of the SOM of rhizosphericsoil might be present as dead microbial residues (necromass),against only 3% as living microorganisms (biomass). More recently,spectrometric and molecular approaches corroborated the essen-tial role of microbial necromass to the maintenance and the accu-mulation of the SOM (Simpson et al., 2007; Clemente et al., 2011;Dumig et al., 2012; Miltner et al., 2012). However, estimations ofthe contribution of fungi and bacteria to SOM stabilization intemperate forest soils remain as scant as the understanding of theirspatial distribution throughout the soil matrix.

Amino sugars (AS) are mostly found in cell walls and extracel-lular polysaccharides of fungi and bacteria, in the exoskeleton of

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139 133

arthropods and in the gut of some worms. The fraction of ASderived from macro and meso organisms is considered negligiblerelatively to that of microorganisms (Amelung, 2001; Simpsonet al., 2004). While 26 AS are identified (Amelung et al., 2008),only glucosamine (GlcN), galactosamine (GalN), muramic acid(MurN) are found in quantifiable concentration and can be used asproxies for their bacterial or fungal origin. Bacterial cell walls arethe unique source of MurN (Glaser et al., 2004; He et al., 2005;Liang et al., 2009). GlcN and GalN have much less specific originsand are both found in considerable amounts in both bacteria andfungi (Amelung et al., 1999; He et al., 2005; Engelking et al., 2007).

AS long-survive their producers and are thus used as proxies formicrobial residues (Amelung, 2001). The introduction of isotopiclabels permits to trace substrate-derived AS, hereafter reported asASS, to capture the response of soil microorganisms to substrateadditions (Boschker and Middelburg, 2002; Glaser, 2005; Amelunget al., 2008). Some studies have focused on compound specificstable isotope techniques using 13C and 15N labels to quantify ASSdynamics (Glaser and Gross, 2005; Liang et al., 2007; Hilscher andKnicker, 2011; He et al., 2011a, 2011b). Until recently, however, 13Canalysis of AS was hampered by undetermined biases on the iso-topic signature of the studied AS resulting from a derivatizationstep for gas chromatography separation (Decock et al., 2009). Toovercome that issue, Bod�e et al. (2009) developed a newmethod for13C analysis of AS by liquid chromatography coupled with isotoperatio mass spectrometry (LC-IRMS) that provides more accurateand precise measurement. This method has been successfully usedto trace ASS during an incubation experiment of 21 days with 13Clabeled plant residues (Bai et al., 2013). This new analytical tool isnow available to determine the effect of the chemical compositionof substrates on the distribution of fungal and bacterial AS residuesthroughout distinct soil compartments.

This study aims at capturing the distribution of newly-producedfungal and bacterial biomasses derived from the added substratesand firmly attached to soil density fractions. We hypothesized thatthe contribution of AS C to the C content of density fraction (frac-tion C) increases with increasing fraction density. We expected themicrobial assimilation of the substrate C among the soil densityfractions to be dictated by substrate availability and thus to revealpreferential microbial habitats. To test these hypotheses, a forestedtopsoil amended with 13C-labeled glycine (readily available to soilmicroorganisms) or finely ground 13C-labeled beech leaves (morecomplex and recalcitrant substrate) were incubated in the labora-tory for 92 days and sequentially separated by density. The for-mation of (13C-labeled) ASS was determined by LC-IRMS (Bod�eet al., 2009). Finally, the formation of fungal and bacterial resi-dues derived from the (13C-labeled) substrates was estimated usingestablished conversion factors.

2. Materials and methods

2.1. Lab incubation

The soil was collected in November 2009 from an even-agedbeech forest of 23-year old at Ebrach, Germany (49�520N,10�270E). Both site and soil have been described in a previous study(Hatton et al., 2012a). Briefly, the soil is an acidic dystric Cambisol(pHH2O: 3.9; sand: 80%; loam: 13%; clay: 7%). The first 2.5 cm of theA-horizon were sampled on 2 m2. The entire volume of soil wasthoroughly mixed and sieved to pass 2 mm. Observable roots andfaunal organisms were removed. The soil was stored for 2 weeksat þ4 �C prior pre-incubation.

The soil was pre-incubated for 15 days at 20 �C (dark; 1:3 soil toair ratio; 42% water holding capacity) prior substrate amendment.Two substrates were used: uniformly 13C-labeled glycine (isotopic

abundance ¼ 98%; SigmaeAldrich, France) and 13C-labeled beechleaves (isotopic abundance ¼ 4.6%). The 13C-labeled leaves wereobtained from 10 year-old beech trees grown in airtight chamber atthe GRAP-CEA Cadarache (France) and 13C-labeled twice a weekusing 13CO2 over a vegetation season (from April to September).Senescent leaves were collected at fall, oven-dried at 25 �C and ballmilled. Glycine and beech leaf fragments were added in an amountcorresponding to the natural annual N input (Hatton et al., 2012a),i.e. 80 mg g�1

soil for glycine and 2.5 mg g�1soil for the leaf fragments

(n ¼ 4 lab replicates).The amended soils were thoroughly mixed by hand and sieved

twice to ensure a homogeneous mixing. Both treatments wereincubated at 20 �C (dark; 1:3 soil to air ratio; 42% water holdingcapacity), aerated once a day for 2 weeks and twice a week untilcollection. The moisture content was readjusted when necessary.The incubation time was set so that the respiration rate, whichincreased after substrate addition, came back to a level comparableto the respiration rate before substrate addition (Bai et al., 2013).The beech leaves treatment was sampled after 92 days. The glycinetreatment was sampled after 7 and 92 days, because the microbialutilization of the glycine C is known to be much faster than forchemically more complex structures (Jones and Murphy, 2007;Rinnan and Baath, 2009). Soils were kept frozen at �20 �C untilanalysis. No macro or meso fauna was detected neither before norafter incubation.

2.2. Sequential density fractionation

Subsamples were sequentially separated by density to isolatestructurally and functionally different soil fractions (Hatton et al.,2012a), shown to be relevant for studying mineral-attached OMdynamics (Moni et al., 2012). The sequential density fractionationwas performed according to Hatton et al. (2012a) with a centrifu-gation kept <2000 g to preserve microbial cells intact. Soils weresuspended in a solution of sodium polytungstate (SPT0, TC-Tungsten Compounds, Germany). For the first separation, the so-lution density was set to 1.65 g cm�3 and centrifuged. The lightfloating fraction was aspirated, rinsed and frozen at �20 �C. Thesame procedure was repeated with solutions of increasing densityto finally obtain five operationally-defined density fractions earlierdescribed by Hatton et al. (2012a): plant debris with little mineralsattached (<1.65 g cm�3), plant aggregates dominated by poorlyaltered plant debris and phyllosilicates minerals(1.65e1.85 g cm�3), microbial aggregates dominated by microbial-processed SOM and phyllosilicates minerals (1.85e2.4 g cm�3) andmineral grains dominated by poorly reactive quartz and feldsparwith little SOM attached (2.4e2.65 g cm�3). The remaining mineralgrains denser than 2.65 g cm�3 were not treated here because ofthe presence of pedogenic oxides, which interfere strongly with thechromatography. However this fraction accounted only for 0.9% and0.2% of soil dry weight and C, respectively.

2.3. Analyses

Carbon content and isotopic composition were determined inthree analytical replicates using an elemental analyzer (CE in-struments, NA 1500 type II) coupled to an isotope ratio massspectrometer (Finnigan, Delta S) at the Technical Platform ofFunctional Ecology (OC 081) at INRA Forest Ecology and Ecophys-iology Unit, INRA Nancy, France.

AS concentration and isotopic composition were determinedusing liquid chromatography coupled to isotope ratio mass spec-trometer (LC-IRMS) at the Isotope Bioscience Laboratory e ISOFYS,Ghent University, Belgium. We followed the method described byBod�e et al. (2009) and two replicates selected from composite

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139134

samples (i.e., from pooled replicates). Samples corresponding to0.3 mg of N were successively hydrolyzed with 6 M HCl (20 ml pergram of sample) at 105 �C for 8 h, filtered through 0.45 mm (GF/C25 mm, Whatman Cat N�1822-025, Germany). Then, AS were pu-rified on a cationic exchange resin (AG50W-X8, 100e200 Mesh,Hydrogen form, Bio-Rad lab.) and eluted by protons prior to LC-IRMS analysis. The chromatographic separation was performedusing a LC pump (Surveyor MS-Pump Plus, Thermo Electron, Bre-men, Germany) mounted with a PA20 CarboPac analytical anion-exchange column (3 � 150 mm, 6.5 mm) and a PA20 guard col-umn (Dionex). Basic AS (GlcN and GalN) were eluted with 2 mMNaOH and a column temperature of 15 �C. The acidic AS (MurN)wasdetermined accordingly to (Bod�e et al., 2013). This method startswith an isocratic part at 10 mMNaOH, after 10 min, 5 mMNaNO3 isadded to the 10 mM NaOH mobile phase to increase the elutionstrength in order to elute the MurN. The eluent flow was set at400 mL min�1 and the column temperature was kept at 15 �C. Theseparated organic compounds were oxidized in 0.03 M Na2S2O8 at99.9 �C, and after acidification with 1.5 M H3PO4 the formed CO2was separated from the liquid phase in the LC Isolink interface(Thermo Electron). Finally, C-isotopic composition of the producedCO2 was analyzed using an isotope ratio mass spectrometer (DEL-TAPLUS XP, Thermo Electron).

2.4. Calculations and statistics

ASS derived from 13C-labeled substrates at time t (ASS,t) wascomputed as follows:

ASS;t ¼ ASt$xð13CÞASt � xð13CÞASt0xð13CÞS � xð13CÞSOM

(1)

with ASt being the amounts of AS at time t and x(13C)S, x(13C)SOM,xð13CÞASt xð13CÞASt0 are the C-isotopic abundance of the initiallyapplied substrate S, of the original SOM and of the AS at time t andt0 (sampled immediately after substrate addition), respectively.xð13CÞASt0 of the leaf incubation was not identical to the originalisotopic composition of the soil AS, due to the presence of 13C-labeled AS of entophytic microorganisms in the plant residues(Bod�e et al., 2013).

Fungal and bacterial C derived from 13C-labeled substrateswere estimated using established conversion factors (Appuhn andJoergensen, 2006; Engelking et al., 2007). As these conversionfactors have been derived from the proportion of AS to microbial Con freshly cultured microorganisms, these factors were only usedfor ASS,t to evaluate the microbial biomass freshly produced fromthe substrates. We did not apply these factors to total AS to esti-mate microbial residues, as have been done elsewhere (vanGroenigen et al., 2010). Fungal C was estimated by multiplying

Table 1Dry weight, C and amino sugar C among soil density fractions isolated from forested topfragments incubated for 92 days and glycine incubated for 7 days and 92 days (n ¼ 4;tosamine (GalN) and muramic acid (MurN) contents are expressed on a C basis (n ¼ 2), peRecovery (%) indicates the recoveries after sequential density fractionation. The descript

Soil fraction Dry weight Fracti

MurN C

g cm�3 Description % mg g�1soil mg g�1 soil mg g�

Bulk soil 100 27.2 ± 0.1 267 ± 59 9.9 ±<1.65 Plant debris 2.6 ± 0.2 8.6 ± 0.1 49.8 ± 3.4 5.9 ±1.65e1.85 Plant aggregates 1.8 ± 0.1 4.9 ± 0.1 36.6 ± 1.7 7.4 ±1.85e2.4 Microbial aggregates 10.2 ± 0.3 9.0 ± 0.4 78.0 ± 3.2 8.9 ±2.4e2.65 Mineral grains 84.9 ± 0.7 2.5 ± 0.1 36.7 ± 0.8 14.5 ±

Recovery (%) 99 ± 1 93 ± 3 75 ± 5 81 ±

the fungal GlcNS,t by 9; fungal GlcNS,t was determined by sub-tracting two times MurNS,t molar concentration from total GlcNS,tas described by Engelking et al. (2007). Bacterial C was estimatedby multiplying the MurNS,t mass concentration by 45. Confidenceintervals were calculated as described by Appuhn and Joergensen(2006).

Comparisons of dry weight, fraction C and substrate C amongsoil fractions were performed using SYSTAT v13 (Systat Software,Inc., Chicago, IL, USA). Differences were tested using general linearmodels followed by a Tukey's post hoc tests. The statistical signif-icance threshold was set at p < 0.05. Correlations with AS and ASSdistributions were obtained using Pearson correlation analysis andreported as R2.

3. Results

3.1. Dry weight, C and AS contents

Recoveries after density fractionation were 99.5 ± 0.4% and91.9 ± 1.4% for dry weight and C, respectively (Table 1). Dry weightand C contents did not differ significantly among the soil fractionsisolated from control and treatments. The only exception was forplant debris fraction isolated from the soil amended with leaves(P < 0.01).

AS concentrations did not differ between control and soilsincubated with beech leaf fragments or glycine (Table 1). AS con-centrations averaged across treatments were 267 ± 59 mgC g�1soil,145 ± 26 mgC g�1soil and 34 ± 15 mgC g�1soil for GlcN, GalN andMurN, representing 0.99 ± 0.22, 0.54 ± 0.10 and 0.13 ± 0.06% of thesoil C, respectively.

AS recoveries after density fractionation did not vary betweentreatments and averaged at 75 ± 5; 69 ± 4 and 58 ± 11% for GlcN,GlaN and MurN, respectively (Table 1). Supplemental Table S1 re-ports the values for individual treatments. GlcN was consistentlythe most abundant AS in all soil fractions, followed by GlaN andMurN. GlcN, GalN and MurN were similarly distributed among thesoil fractions regardless to the treatments (Fig. 1). 24 ± 5% of therecovered AS were found in plant debris (<1.65 g cm�3), 18 ± 3% inplant aggregates (1.65e1.85 g cm�3), 40 ± 5% in microbial aggre-gates (1.85e2.4 g cm�3) and 19 ± 3% in mineral grains(2.4e2.65 g cm�3). The AS C to fraction C ratio increased consis-tently with soil fraction density for all three AS.

3.2. Substrate-derived C

87.8 ± 3.8% of the beech leaf C initially applied (leaf CS,0) wasrecovered in soils after 92 days of lab incubation (Table 2). Re-covery after density fractionation was 96 ± 6% of the residual leafC found after 92 days (leaf CS,92d). Leaf CS,92d, was unequally

soil. Values are means ± standard errors from a control and three treatments: leafTable S1 displays the values for individual treatments). Glucosamine (GlcN), galac-r gram of soil and per gram of fraction C (i.e, the amount of C of a given soil fraction).ion of the soil density fractions is reprinted from Hatton et al. (2012a).

on C GlcN C GalN C

1 fraction C mg g�1 soil mg g�1 fraction C mg g�1 soil mg g�1 fraction C

2.2 145 ± 26 5.4 ± 1.0 33.8 ± 14.9 1.3 ± 0.60.4 23.1 ± 1.4 2.7 ± 0.2 3.7 ± 0.6 0.4 ± 0.10.3 17.5 ± 0.8 3.6 ± 0.2 2.9 ± 0.9 0.6 ± 0.20.4 41.3 ± 2.0 4.7 ± 0.2 7.3 ± 2.0 0.8 ± 0.20.5 17.8 ± 0.3 7.0 ± 0.2 5.5 ± 1.9 2.2 ± 0.7

5 69 ± 4 74 ± 4 58 ± 11 63 ± 12

Fig. 1. Mass distribution of total amino sugars (AS) and amino sugars formed after atime t from (glycine ASS,t; t ¼ 7 or 92 days) or beech leaf fragments (leaf ASS,t; t ¼ 92days) among soil density fractions isolated from a forested topsoil (n ¼ 2). All distri-butions are expressed as percent of what was recovered after sequential densityfractionation. Error bars are standard errors. Triangles show the distribution of thetotal C among soil density fractions (fraction C). White circles represent the residualleaf-derived C recovered after 92 days (leaf CS,92d). White squares represent the re-sidual glycine-derived C recovered after 7 days (glycine CS,7d). Dark squares representthe residual glycine-derived C recovered after 92 days (glycine CS,92d). Dark barsrepresent the recovered leaf ASS,92d. Gray bars represent the recovered glycine ASS,7d(left) and glycine ASS,92d (right). White bars show the recovered AS C. All durationsrefer to days of lab incubation (20 �C, dark, 42% water holding capacity). Thedescription of the soil density fractions is reprinted from Hatton et al. (2012a).

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139 135

distributed among the soil density fractions (Fig. 1): 56 ± 7% ofthe recovered leaf CS,92d was found in plant debris(<1.65 g cm�3), 22 ± 2% in plant aggregates (1.65e1.85 g cm�3),16 ± 4% in microbial aggregates (1.85e2.4 g cm�3) and 6 ± 1% inmineral grains (2.4e2.65 g cm�3).

8.93 ± 0.14% and 5.44 ± 0.11% of the glycine C initially applied(glycine CS,0) were recovered in soils after 7 and 92 days of lab in-cubation, respectively (Table 2). Recoveries after density fraction-ation increased from 59 ± 4% to 76 ± 4% of the residual glycine Cafter 7 days (glycine CS,7d) and 92 days (glycine CS,92d), respectively.Glycine CS,7d and glycine CS,92d were similarly distributed among

Table 2Substrate-derived C and amino sugars C derived from glycine or beech leaf fragments(substrate C) is expressed as percent of applied (n¼ 4). Glucosamine C, galactosamine C anC, GalNS C and MurNS C, respectively (n ¼ 2). GlcNS, GalNS and MurNS are expressed relativMurN C contents (reported in Table S1). The description of the soil density fractionsmeans ± standard errors.

Treatment Soil fraction Substrate C GlcNS C

Days g cm�3 Description % applied mg g�1CS,0

Beech leaves 92 Bulk soil 87.8 ± 3.8 1729 ± 17<1.65 Plant debris 47.7 ± 4.7 420 ± 701.65e1.85 Plant aggregates 18.2 ± 1.0 151 ± 121.85e2.4 Microbial aggregates 13.5 ± 3.3 182 ± 182.4e2.65 Mineral grains 5.2 ± 0.3 86 ± 8

Recovery (%) 85 ± 6 49 ± 7

Glycine 7 Bulk soil 8.93 ± 0.14 975 ± 21<1.65 Plant debris 0.98 ± 0.04 109 ± 261.65e1.85 Plant aggregates 1.11 ± 0.08 141 ± 271.85e2.4 Microbial aggregates 2.37 ± 0.18 342 ± 892.4e2.65 Mineral grains 0.79 ± 0.24 158 ± 24

Recovery (%) 59 ± 4 77 ± 20

92 Bulk soil 5.44 ± 0.11 942 ± 25<1.65 Plant debris 0.79 ± 0.02 117 ± 401.65e1.85 Plant aggregates 0.88 ± 0.04 130 ± 331.85e2.4 Microbial aggregates 1.92 ± 0.13 428 ± 532.4e2.65 Mineral grains 0.56 ± 0.18 137 ± 13

Recovery (%) 76 ± 4 86 ± 25

soil density fractions (Fig. 1). In average, 19 ± 1% of the recoveredglycine CS,t was found in plant debris (<1.65 g cm�3), 21 ± 1% inplant aggregates (1.85e2.4 g cm�3), 46 ± 3% inmicrobial aggregates(1.85e2.4 g cm�3) and 14 ± 3% in mineral grains (2.4e2.65 g cm�3).

3.3. Substrate-derived ASS

Leaf-derived ASS produced after 92 days (leaf ASS,92d) are re-ported in Table 2. GlcNS was consistently the most abundant ASS,followed by MurNS and GalNS. Leaf GlcNS,92d, GalNS,92d andMurNS,92d averaged at 7.1 ± 1.1; 1.4 ± 0.2 and 14 ± 5 mg g�1 of GlcN,GalN and MurN, respectively. Recoveries after density fractionationwere of 49 ± 7%, 94 ± 17% and 51 ± 17% for GlcNS, GalNS andMurNS,respectively (Table 2). Leaf ASS,92d were unequally distributedamong the soil density fractions (Fig. 1). Leaf ASS,92d were mostabundant in the plant debris fraction (<1.65 g cm�3), with 50 ± 9%,62 ± 13% and 38 ± 11% of the recovered leaf GlcNS,92d, GalNS,92d andMurNS,92d, respectively. The rest of the recovered leaf ASS,t wassimilarly distributed among plant aggregates (1.65e1.85 g cm�3),microbial aggregates (1.85e2.4 g cm�3) and mineral grains(2.4e2.65 g cm�3), regardless ASS type.

Glycine-derived GlcNS, GalNS and MurNS produced after 7 days(glycine ASS,7d) and 92 days (glycine ASS,92d) were similarlydistributed among the soil fractions (Table 2), suggesting that mostglycine ASSwere produced in less than 7 days after glycine addition.Glycine ASS,7d and glycine ASS,92d averaged at 77 ± 9 ngC g�1soil,9 ± 2 ngC g�1soil and 3 ± 1 ngC g�1soil for GlcNS, GalNS and MurNS,respectively. Glycine ASS recoveries after density fractionationweresimilar for glycine ASS,7d and glycine ASS,92d and averaged at82 ± 16, 60 ± 16 and 77 ± 21% for GlcNS, GalNS and MurNS,respectively. Glycine ASS,7d and glycine ASS,92d were similarlydistributed among the soil density fractions (Fig. 1). In average,glycine ASS,t all peaked in microbial aggregates (1.85e2.4 g cm�3)that accounted for 49 ± 7, 48 ± 13 and 57 ± 19% of the glycine GlcNS,GalNS and MurNS recovered after density fractionation (Fig. 1). Therest of the glycine ASS,t recovered after density fraction was simi-larly distributed among plant debris (<1.65 g cm�3), plant aggre-gates (1.65e1.85 g cm�3) and mineral grains (2.4e2.65 g cm�3),regardless to the ASS type.

among soil density fractions isolated from a forested topsoil. Substrate-derived Cd muramic acid C derived from (13C-labeled) glycine or beech leaves are noted GlcNS

ely to the substrate C initially applied (CS,0) and relatively to the total GalN, GlcN andis from Hatton et al. (2012a). n.a. means non applicable. Data are reported as

GalNS C MurNS C

mg g�1 GlcN C mg g�1CS,0 mg g�1 GlcN C mg g�1CS,0 mg g�1 GlcN C

9 7.1 ± 1.1 185 ± 26 1.4 ± 0.2 418 ± 95 14 ± 58.8 ± 1.6 107 ± 19 4.7 ± 0.9 82 ± 14 26 ± 115.0 ± 0.6 21 ± 2 1.4 ± 0.2 51 ± 33 30 ± 252.4 ± 0.3 28 ± 4 0.7 ± 0.1 46 ± 29 7 ± 62.4 ± 0.2 18 ± 3 1.1 ± 0.2 34 ± 17 6 ± 3n.a. 94 ± 17 n.a. 51 ± 17 n.a.

2 0.29 ± 0.05 106 ± 36 0.058 ± 0.011 26.6 ± 5.0 0.07 ± 0.020.16 ± 0.02 12 ± 3 0.039 ± 0.005 2.5 ± 0.5 0.06 ± 0.020.34 ± 0.04 9 ± 3 0.041 ± 0.008 3.1 ± 1.4 0.13 ± 0.090.33 ± 0.05 32 ± 11 0.056 ± 0.011 12.4 ± 6.2 0.13 ± 0.110.32 ± 0.03 9 ± 1 0.038 ± 0.003 4.3 ± 3.8 0.06 ± 0.05n.a. 58 ± 22 n.a. 84 ± 32 n.a.

4 0.28 ± 0.05 105 ± 36 0.058 ± 0.011 40.3 ± 11.5 0.10 ± 0.040.18 ± 0.03 15 ± 8 0.048 ± 0.013 4.1 ± 2.2 0.09 ± 0.060.31 ± 0.05 9 ± 4 0.044 ± 0.010 3.7 ± 3.3 0.15 ± 0.160.41 ± 0.03 28 ± 8 0.050 ± 0.008 16.2 ± 4.9 0.17 ± 0.130.28 ± 0.02 12 ± 2 0.049 ± 0.004 4.2 ± 3.7 0.05 ± 0.05n.a. 61 ± 24 n.a. 70 ± 27 n.a.

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139136

3.4. Estimated substrate-derived bacterial and fungal C

Fig. 2 illustrates the estimated contribution of leaf and glycine-derived bacterial and fungal C to CS,t among soil fractions. Theoverall microbial C contribution to CS,t was lesser for leaves (6e13%)than for glycine, and for glycine after 7 days (23e32%) than forglycine after 92 days (37e56%). For the soils amended with leaves,these differences were driven by the substrate-derived bacterial C,with C to CS,t ratio of 2.6e3.3%. Conversely, these differences weredriven by the substrate-derived fungal C for the soils amendedwithglycine, with a fungal C to CS,t ratios of 21e26% and 32e40% after 7and 92 days (Fig. 2). Leaf and glycine derived bacterial and fungal Crelative to CS,t both tended to increase with increasing fractiondensity. Glycine derived bacterial and fungal C relative to CS,t ten-ded to be greater after 92 days than after 7 days in all soil densityfractions. The ratios of the substrate derived fungal to bacterial Cwere consistently greater for glycine than for leaves. The ratios ofthe substrate derived fungal to bacterial C did not vary amongdensity fractions for both glycine and leaves.

4. Discussion

4.1. Microbial C contribution to fraction C

More than 90% of the soil AS are present in residues of deadmicrobial cells (Amelung, 2001; Amelung et al., 2008), so that AScan be used to assess the spatial distribution of microbial residuesin soils. Here, AS accounted for 1.7 ± 0.1% of soil C, what is com-parable with previous studies reviewed by Glaser (2005). The GlcNC to MurN C ratio (8 ± 3) was at the low end of what was found inliterature (Joergensen and Wichern, 2008). This relatively high

Fig. 2. Estimated contributions of bacterial and fungal C residues to residual substrateC (CS,t) among soil density fractions sequentially isolated from a forested topsoil(n ¼ 2). The bacterial and fungal C contributions are expressed as percent of fractionCS,t (lower panel) and relatively as fungal C to bacterial C ratios (upper panel). Whitebars represent the microbial C formed from beech leaves after 92 days of incubation(leaf microbial C). Gray bars represent the microbial C formed from glycine (glycinemicrobial C) after 7 days (dark gray) and 92 days (light gray). Errors bars represent the95% confidence limits of the factors suggested by Appuhn and Joergensen (2006) forthe conversion of the substrate-derived muramic acid or fungal glucosamine intobacterial (without pattern) or fungal (with pattern) biomasses, respectively. The soilfractions were isolated as described by Hatton et al. (2012a) and described in Tables 1and 2.

MurN contribution and the low C/N ratio (5.5) of the microbialbiomass (Zeller and Dambrine, 2011) are indicative of a microbialbiomass dominated by bacteria. This might also be attributed to thepreferential stabilization of MurN with sandy soils hypothesized byDecock et al. (2009).

The AS C to fraction C ratios reported in Table 1 suggest agreater contribution of microbial C residues to the mineral-attached C of the microbial aggregates fraction(1.85e2.4 g cm�3) and, mostly, of the mineral grain fraction(2.4e2.65 g cm�3). This view is supported by earlier observationsamong other soil density fractions (Sollins et al., 2006, 2009;Bonnard et al., 2012; Hatton et al., 2012a) and otheroperationally-defined soil fractions (von Lützow et al., 2007;Grandy and Neff, 2008). This finding might be partly attributedto the progressive accumulation of microbial residues along withthe progressive transfer of litter-derived materials from plantdebris (<1.65 g cm�3) to plant aggregates (1.65e1.85 g cm�3) andmicrobial aggregates (1.85e2.4 g cm�3) described by Hatton et al.(2012a). Another possible enon-exclusivee explanation is thepreferential retention of microbial C residues through chemi-sorption with phyllosilicates minerals dominating microbial ag-gregates (1.85e2.4 g cm�3). Unlike for the microbial aggregates,the mineral grain fraction (2.4e2.65 g cm�3) is dominated bypoorly reactive quartz and feldspar minerals (Hatton et al., 2012a)so that sorption mechanisms of microbial C residues are ratherlimited. We posit that the high contribution of microbial residuesrelatively to the mineral-attached SOM of the mineral grainsfraction (2.4e2.65 g cm�3) indicates a preferential habitat for soilmicroorganisms. Moreover, these results together with the longermean residence times measured in similar soil density fractions(Sollins et al., 2006, 2009; McFarlane et al., 2013) support the viewrecently made by Miltner et al. (2012) that microbial residues arean important source of stable SOM. Finally, the relatively higherrenewal rate of MurN compared to GlaN and GlcN (Table 2) tend tosupport the view that bacterial C residues do not last as long asfungal C ones in soils (Six et al., 2006).

4.2. Microbial C from glycine and beech leaves among soil fractions

Freshly produced ASS derived from glycine and beech leavesinform on bacterial and fungal response to the applied substrates.Our results showed that the production of ASS is both substrate andAS specific as earlier described by He et al. (2011a) and Bai et al.(2013). Here, we investigate the microbial C assimilation fromready available substrate (glycine) and from the substrate naturallypresent on site (beech leaves) among soil density fractions, i.e. formicrobial components firmly attached to soil assemblages.

Substrate use efficiency was estimated from the amount ofsubstrate derived microbial C relative to the amount of substrateutilized by the microorganisms (microbial-C þ amount mineral-ized). Glycine appeared to be much less efficiently used for mi-crobial growth (2.5 ± 0.3% after 7 and 92 days) than beech leaves(36 ± 10% after 92 days). Similarly low substrate use efficiency uponglycine was reported by Sugai and Henrichs (1992), with ca. 60% ofglycine C mineralized in 2 h only. Glycine is rapidly assimilated bythe soil microorganisms (Webster et al., 1997; Henry and Jefferies,2003; Jones and Murphy, 2007) and further allocated to a varietyof compounds, such as sugars (Derrien et al., 2007), enzymes(Hatton et al., submitted for publication), etc. Here, we posit thatthe microbial recycling of part of the primary microbial productscan result in large C losses that could partly explain the apparentlow microbial use efficiency of glycine.

We found that 35 and 49% of the glycine C recovered among soildensity fraction after 7 and 92 days was of microbial origin,respectively (Fig. 2). Given the rapid turnover of glycine in soils

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139 137

(Henry and Jefferies, 2003; Jones and Murphy, 2007), one mayexpect that the large majority of the glycine C attached to soildensity fractions should be part of the microbial C after 7 and 92days of incubation. In fact, microbial activities explained 90% of theglycine C recovered among soil density fractions isolated after only8 h under the same experimental conditions (Hatton et al.,submitted for publication). The conversion factors used for Fig. 2are rightfully disputable as they have been derived from a limitednumber of bacterial and fungal cultures (Appuhn and Joergensen,2006; Engelking et al., 2007), which might not be representativefor the soil microbial communities. We further posit that thisapparent underestimation is also due to the differences in matrix ofculture conditions and soil. While the secretion of extracellularmicrobial products in cultures is limited due to the larges abun-dance of resources, microorganisms inhabiting in soils most likelyallocate a larger fraction of the assimilates to the production ofextracellular products (Schimel and Schaeffer, 2012; Burns et al.,2013), such as extracellular exopolysaccharides and diffusive en-zymes (Hatton et al., submitted for publication). Therefore, themicrobial C assessed using the ASS is rather an estimation of themicrobial cell C than the total microbial C.

The low extent of glycine assimilation by bacteria was unex-pected given that glycine is well known to be rapidly assimilated bysoil microorganisms (Jones and Murphy, 2007; Rinnan and Baath,2009). The little accumulation of glycine C in bacterial AS mightbe explained by (1) a preferential uptake of glycine C by gramnegative bacteria (G�), known to have ~20% less AS than Gþ bac-teria (Engelking et al., 2007) and to dominate the initial stage ofdecomposition (Rinnan and Baath, 2009; Fontaine et al., 2011;Herman et al., 2012). This would underestimate the contributionof the bacterial community to the incorporation of glycine. More-over, the little amount of added glycine (80 mg g�1

soil) necessarilyresults in (2) a physically limited substrate availability for bacteriamore than for Fungi. Such small amount of glycine addition arespatially highly heterogeneously distributed at scales relevant tomicroorganisms (Hatton et al., submitted for publication; Hattonet al., 2012b), what could indirectly favor fungal assimilation. Un-like bacteria, fungi benefits from hyphen high prospection capa-bilities that confer a considerable competitive advantage overbacteria, assimilating a substrate as limiting as the glycine appliedin this study. Fungi would likely exhaust the limited glycineresource before bacteria have had the opportunity to access thesubstrate, e.g. through the “fungal highway” (Kohlmeier et al.,2005).

As earlier pointed out in a literature review by Strickland andRousk (2010), our results do not support the view that funginecessarily dominate the decomposition of plant litter (de Boeret al., 2005). The lower GlcNS to MurNS ratios found for leaf ASS(4.1 ± 0.3; Table 2) compared with the original GlcN to MurN ratio(8.3 ± 0.3, Table 1) could be attributed to the fact that bacteria drivethe initial stage of litter decomposition (Bernard et al., 2007),presumably through the mineralization of easily available C forms(Poll et al., 2008). A similar observation was recently made by Bod�eet al. (2013), in a study that also pointed the necessary coupling e

without reciprocitye of bacterial activities to fungal activities at theearly stage of litter decomposition in soils. In the later stages ofdegradation, more specialized saprotrophic fungi take over andassimilate the more recalcitrant compounds left behind, such aslignin and cellulose (de Boer et al., 2005; Fontaine et al., 2011).However, this difference in GlcN toMurN ratios might also build upovertime through the better preservation of fungal residuescompared to bacterial ones (Six et al., 2006). The relatively highrenewal rate of the MurN compared to GlaN and GlcN (Table 2)further support the view that bacterial residues do not last as longas fungal ones in soils.

The lowamount of MurN and leaf MurNS recovered after densityfractionation (58 ± 11 and 51 ± 17%, respectively) suggest that alarge fraction of bacterial bio- and necro-masses that is either freeand/or poorly attached to the soil is similarly lost during thesequential density fractionation of soils. This view is supported bythe amount of GlcNS versus GlcN recovered after density fraction-ation (Tables 1 and 2). When fungal GlcNS dominated the overallGlcNS pool, like for glycine, the amount of GlcNS and (total) GlcNrecovered after density fractionation was comparable. Conversely,less GlcNS was recovered after density fractionation than (total)GlcN when bacterial GlcNS dominated the overall GlcNS pool, likefor beech leaves with 49 ± 7% of the leaf GlcNS recovered versus75 ± 5% for (total) GlcN. We posit that the proportion of bacterialversus fungal GlcNS determines the extent to which GlcNS isrecovered after density fractionation. The greater recovery of fungalGlcNS was attributed to the intimate association of fungal hyphaewith soil assemblages, while the lesser recovery of bacterial GlcNSwas attributed to the fraction of bacterial cell residues that is free orloosely attached to the soil. Finally, the greater recovery of glycineMurNS,t (77 ± 21%) compared with leaf MurNS,92d (51 ± 17%) canprobably be attributed to the fact that most bacteriawho had accessto the glycine were probably associated to fungi, making them lesslikely to be lost during the fractionation procedure.

4.3. Microbial distribution and ecological implications

Unlike leaf fragments, glycine is a readily available substrate tosoil microorganisms (Jones and Murphy, 2007; Rinnan and Baath,2009). While leaf ASS distribution among soil density fractionswas used to inform on the spatial distribution of naturally occurringmicrobial processes of litter degradation (i.e. during the first 92days of litter decay), glycine ASS distribution was used to revealwhere the microbial activity is preferentially located. The com-parison of the two distributions was further used to refine ourunderstanding of the functioning of the soil system during SOMdegradation and formation.

Fig. 1 illustrates a strong correlation between the distribution ofglycine ASS,t C and the residual glycine CS,t (R2 > 0.91), both peakingin the microbial aggregates fraction dominated by phyllosilicatesminerals (1.85e2.4 g cm�3). This was attributed to the fact that themajority of the residual glycine CS,t is most likely found as microbialC residues (Fig. 2). The fact that the glycine ASS,t C peaks in themicrobial aggregates fraction (1.85e2.4 g cm�3) further suggeststhat the microbial activity efungal but also bacteriale is prefer-entially located there. Fig. 1 also illustrates a very good correlationbetween leaf CS,t and leaf ASS,92d (R2 ¼ 0.97), both peaking in plantdebris (<1.65 g cm�3). As earlier reported by Foster (1988) andYoung et al. (2008), these results indicate that the microbial C ismostly produced where the resource is located.

The AS distribution is indicative of the microbial necromass(Amelung et al., 2008) and reflects the integrated state of the mi-crobial residues produced over time. The fact that AS distributesimilarly to glycine ASS,t (R2 � 0.87) indicates that the microbialaggregates fraction dominated by phyllosilicates (1.85e2.4 g cm�3)not only accumulates microbial necromasses, but also constitutes apreferential habitat for living biomass. The comparison with whatwas obtained for the beech leaves further suggests that, in naturallitter decomposition processes, the microbial residues do notremain in plant debris (<1.65 g cm�3) where they are principallyproduced (Fig. 2), but moves towards the microbial aggregates(1.85e2.4 g cm�3). Such movement might be explained by thedynamic transfer of litter-derived materials from plant debris(1.65 g cm�3) to microbial aggregates (1.85e2.4 g cm�3) along withOM decay (Hatton et al., 2012a).

P.-J. Hatton et al. / Soil Biology & Biochemistry 79 (2014) 132e139138

Finally, our results show that the contribution of microbial C tothe residual substrate Cs,t increases with increasing soil fractiondensity both for glycine and beech leaf fragments (Table 2; Fig. 2).The same pattern was observed for the contribution of AS C to thefraction C, i.e. also it increases with fraction density (Table 1). Thesefindings reinforce the hypothesis that the accumulation of themicrobial products along with density might also be attributed to apreferential stabilization. Possible explanations are (1) increasingstabilization efficiency of microbial products, from plant debris toaggregates and, finally, mineral grains and (2) specific physico-chemical interactions with the soil matrix (Chenu and Stotzky,2002). AS have been shown to adsorb to some crystalline mineralforms (Kaiser and Zech, 2000), which dominate the mineral phaseof plant and microbial aggregates (Hatton et al., 2012a), and poorlycrystalline (hydr)oxides (Mikutta et al., 2010; Keiluweit et al., 2012),principally found in the mineral grains isolated between 2.4 and2.65 g cm�3 (Hatton et al., 2012a). Our findings may also beattributed to a segregation of functionally different microbialcommunities among the different soil compartments: while themicroorganisms feeding on plant litters would be preferentiallyinhabiting low-density fractions, the microorganisms inhabitinghigh-density fractions would preferentially feed on microbial res-idues and/or assimilate labile leaf components, which would bemore efficiently assimilated and stabilized by soil microorganisms(Cotrufo et al., 2013). The hypothesis that functionally differentmicrobial communities are differently distributed throughout thesoil matrix is supported by earlier studies investigating differentaggregate-size fractions (Dorodnikov et al., 2009; Davinic et al.,2012). Our research, however, did not specifically address this hy-pothesis, which would benefit from additional research.

5. Conclusions

This study uses compound specific stable isotope analysis ofglucosamine, galactosamine and muramic acid to assess the dis-tribution of microbial residues derived from two contrasting sub-strates, beech leaves and glycine, and gain insight into the spatialdistribution of microbial residues among soil fractions separated bydensity. We showed that:

1) After 12 weeks of incubation, the microbial assimilation ofbeech leaf C appears slightly dominated by bacteria, while fungiclearly dominate the microbial assimilation of glycine C.

2) The contribution of the fungal and bacterial AS C to the mineral-attached C tends to increase with fraction density. Similarly, thecontribution of the glycine and leaf-derived AS C to the residualsubstrate C also tend to increase with fraction density. Thissuggests an increasing stabilization efficiency of the microbial Cresidues along with increasing fraction density, from plantdebris to aggregates and, finally, mineral grains.

3) The microbial assimilation of the substrate C throughout the soilmatrix appeared to be initially governed by the accessibility ofthe substrate. With time, however, the distribution of microbialC residues appear to be driven by a progressive SOM transfer,from plant debris to microbial aggregates.

Acknowledgments

The Institut National de la Recherche Agronomique (INRA-EFPA), the R�egion Lorraine (grants #12000292 and #12000184), theInternational Atomic Energy Agency (IAEA, CRP D12011) and theESF-Molter programme have financially supported this work (grant#2961). We are in debt to Christian Hossan (PTEF, INRA) for EA-IRMS analyses. The UMR 1137 and the UR 1138 are supported bythe French National Research Agency through the Laboratory of

Excellence ARBRE (ANR-12-LABXARBRE-01). We thank the re-viewers for their constructive comments that greatly enhanced themanuscript.

Appendix A. Supplementary data

Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.soilbio.2014.09.013.

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