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Soil Biology & Biochemistry 42 (2010) 1009e1016

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Soil Biology & Biochemistry

journal homepage: www.elsevier .com/locate/soi lb io

Carbon input belowground is the major C flux contributing to leaf littermass loss: Evidences from a 13C labelled-leaf litter experiment

Rubino M.a,*,1, Dungait J.A.J.b,2, Evershed R.P.b, Bertolini T.a, De Angelis P.c, D’Onofrio A.a,Lagomarsino A.c, Lubritto C.a, Merola A.a, Terrasi F.a, Cotrufo M.F.a,3

aDipartimento di Scienze Ambientali, Seconda Università di Napoli, via Vivaldi, 43, 81110 Caserta, ItalybOrganic Geochemistry Unit, Bristol Biogeochemistry Research Centre, School of Chemistry, University of Bristol, Cantock’s Close, Bristol BS8 1TS, UKcDipartimento di Scienze dell’Ambiente Forestale e delle sue Risorse, Università degli studi della Tuscia, Via S.Camillo de Lellis, 01100 Viterbo, Italy

a r t i c l e i n f o

Article history:Received 30 June 2009Received in revised form18 February 2010Accepted 22 February 2010Available online 26 March 2010

Keywords:Stable carbon isotopesLeaf litter decompositionSoil organic matterSoil microbial biomassCarbon cyclingPLFA

* Corresponding author at: Division of Marine and107-121 Station Street, 3195 Aspendale, Victoria, Austrfax: þ61(0) 392394444.

E-mail address: mauro.rubino@csiro.au (M. Rubino1 Present address: Centre for Ice and Climate, Niels

Copenhagen, Juliane Maries Vej 30, 2100 Copenhagen2 Present address: Biogeochemistry of Soils & Wate

gramme for Sustainable Soil Function (Soil-CIP), NorthDevon EX20 2SB, UK.

3 Present address: Department of Soil and CroUniversity, 80523 Fort Collins, Colorado, USA.

0038-0717/$ e see front matter � 2010 Elsevier Ltd.doi:10.1016/j.soilbio.2010.02.018

a b s t r a c t

Partitioning of the quantities of C lost by leaf litter through decomposition into (i) CO2 efflux to theatmosphere and (ii) C input to soil organic matter (SOM) is essential in order to develop a deeperunderstanding of the litter-soil biogeochemical continuum. However, this is a challenging task due to theoccurrence of many different processes contributing to litter biomass loss. With the aim of quantifyingdifferent fluxes of C lost by leaf litter decomposition, a field experiment was performed at a short rotationcoppice poplar plantation in central Italy. Populus nigra leaf litter, enriched in 13C (d13C w þ160&) wasplaced within collars to decompose in direct contact with the soil (d13C w �26&) for 11 months. CO2

efflux fromwithin the collars and its isotopic composition were determined at monthly intervals. After 11months, remaining litter and soil profiles (0e20 cm) were sampled and analysed for their total C and 13Ccontent. Gas chromatography (GC), GCemass spectrometry (MS) and GC-combustion-isotope ratio(GC/C/IRMS) were used to analyse phospholipid fatty acids (PLFA) extracted from soil samples to identifythe groups of soil micro-organisms that had incorporated litter-derived C and to determine the quantityof C incorporated by the soil microbial biomass (SMB). By the end of the experiment, the litter had lostabout 80% of its original weight. The fraction of litter C lost as an input into the soil (67 � 12% of the totalC loss) was found to be twice as much as the fraction released as CO2 to the atmosphere (30 � 3%), thusdemonstrating the importance of quantifying litter-derived C input to soils, in litter decompositionstudies. The mean d13C values of PLFAs in soil (d13C ¼ �12.5&) showed sustained incorporation of litter-derived C after one year (7.8 � 1.6% of total PLFA-C). Thus, through the application of stable 13C isotopeanalyses, we have quantified two major C fluxes contributing to litter decomposition, at macroscopic andmicroscopic levels.

� 2010 Elsevier Ltd. All rights reserved.

1. Introduction

Leaf litter decomposition is a key process in vegetated terrestrialecosystems, contributing to the formation of soil organic matter(SOM, Liski et al., 2002) as well as to ecosystem respiration. At the

Atmospheric Science, CSIRO,alia. Tel.: þ61(0) 392394634;

).Bohr Institute, University of, Denmark.r Group, Cross Institute Pro-Wyke Research, Okehampton,

p Sciences, Colorado State

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soil surface, litter is fragmented by biotic agents, e.g. macro-invertebrates, and abiotic processes, such as sunlight, wind and rain(Cortez, 1998; Cortez and Bouché, 1998; Dilly et al., 2004; Schadlerand Brandl, 2005), allowing subsequent physical movement intothe soil profile by leaching. The activities of micro-organisms(bacteria and fungi), both within surface litter and after incorpo-ration into the soil profile, further decomposes the litter. Theircatabolic activity ultimately mineralises organic matter to CO2,though microbial metabolites are also produced that contributedirectly to SOM formation (Rubino et al., 2007). The quantificationof these major C fluxes contributing to litter decomposition has notyet been applied to the same sample due to the intrinsic complexityof the decomposition process and the limitations of the prevailingtechnical approach, i.e. litterbags (Bocock and Gilbert, 1957), widelyused to measure litter mass loss and calculate litter decay rates(Gholz et al., 2000).

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e10161010

Litterbags are useful in decomposition experiments as theyconfine the litter and prevent contamination from new litter inputs(Adair et al., 2008; Chadwick et al., 1998; Smith and Bradford, 2003;Yamashita and Takeda, 1998). However, they can fundamentallyalter the decomposition environment by influencing naturalinteractions with soil macrofauna, for instance (Smith andBradford, 2003; St. John, 1980), an effect demonstrated to bedependent on bag mesh size (Bradford et al., 2002; St. John, 1980).At the same time, litterbags can prevent fragmented litter fromphysical transport into the underlying soil horizons, and, thereafterincorporation into SOM. Under such conditions, the amount ofC lost during litter decomposition in studies using litterbags isusually assumed to be caused by microbial respiration in situ,whereas the potential for C to enter the soil and to become incor-porated into SOM is rarely investigated. Thus, in this study, we haveapplied an alternative technique, using the method described bySubke et al. (2004).

During its decomposition, leaf litter loses biomass. Two majorC fluxes are involved: (i) CO2 released to the atmosphere and (ii)organic compounds translocated into the soil. Measuring litterC fluxes is challenging but advances in stable IRMS have enabledC inputs to be differentiated from existing SOM, thereby allowingmore accurate assessments of C pools in soils. For example, Subkeet al. (2004) used 13C-labelled Spruce litter, applied within collarsto the soil of a forest stand, to partition soil respiration and quantifythe contribution of litter decomposition to soil CO2 efflux. Theadvantage of using stable isotopes is that the environment underinvestigation remains relatively unaltered. By substituting thenative leaf litter with a similar, but isotopically labelled litter, theinput of C in the soil from the decomposing litter can be determinedas a variation in the 13C isotope in SOM. Therefore, the change inisotopic composition is proportional to the amount of C derivedfrom the litter and this amount can be quantified applying a simpletwo end members mixing model (Balesdent et al., 1987). Severalstudies have used this approach to quantify the C input fromdecaying litter material to SOM (Bird et al., 2008, 2003; Rasmussenet al., 2008).

The same approach can be used to determine the input of C intothe soil microbial biomass (SMB, Evershed et al., 2006). Gas chro-matographic (GC) analyses of individual fatty acids derived frommicrobial membrane phospholipid fatty acids (PLFA), allowscalculation of abundance of biomarker compounds which can beused to investigate the dynamics of microbial populations in soils(Dungait et al., 2005). Thereby, the introduction of 13C-labelledsubstrates to soils followed by GC-IRMS analyses of individual PLFAprovides both quantitative and chemotaxonomic information forgroups of micro-organisms utilizing a given substrate. For example,the application of 13C-PLFA analysis has suggested that elevated CO2increased fungal activity and microbial metabolism of cellobiose(Phillips et al., 2002).

Described herein are the results of an investigation aimed atquantifying C fluxes from leaf litter during decomposition. Toachieve this, we incubated 13C-labelled litter in direct contactwith the soil and thereafter determined the change of soil d13Cvalues as well as the quantity and d13C values of the respired CO2.Using 13C-PLFA analysis we were able to (i) identify the groups ofmicro-organisms that had incorporated litter-derived C below-ground and (ii) quantify the amount of litter-derived C incorpo-rated into the SMB. To our knowledge, this is the first litterdecomposition study where mass loss is budgeted in terms ofCO2 efflux to the atmosphere and C input belowground, and alsorelated directly to the microbial community undertaking thedecomposition. This represents an important step towardsa deeper and mechanistic comprehension of the litter-soilbiogeochemical continuum.

2. Materials and methods

2.1. Study site and experimental setup

The experimental site is located in a 9-ha short rotation coppiceculture of poplars (Populus x euramericana) in Central Italy (Tuscania;42�220 N,11�480 E, altitude 150 m). The soil is classified, according tothe USDA system, as Pachic Xerumbrempts, with a deep A horizon,and an O horizon of 2e3 cm, generally distinct as a litter anda fermentation layer. The soil is loam/silt-loam and the pH range is4.8e5.5. For further information on soil physical and chemicalproperties see Hoosbeek et al. (2004).

This was the site of a previous FACE experiment (ScarasciaMugnozza et al., 2000) which had been in place for 6 years, andwas used to take advantage of the experimental design, i.e.: 6 largeexperimental plots (314 m2) with 2 continuing nitrogen fertiliza-tion treatments (Calfapietra et al., 2007). Soil temperature wascontinuously measured at the site, as described by Miglietta et al.(2001). The effect of the previous FACE experiment in terms ofambient or increased concentrations of CO2 did not significantly(P< 0.01) affect subsequent d13C values during this experiment (seebelow).

The 13C-labelled litter was obtained from Populus nigra saplingsgrown for two months in large growth chambers (E.T.G., Florence,Italy), at ambient CO2 concentration, under continuous label with13CO2 (Spectragas, 99% 13CO2). The amount of the injected purelabelled CO2 was adjusted weekly, according to the growth of theplants. The gas flow rates were in the range 0.1e1 ml min�1.The temperature/light cycle was 14 h 25 �C/light on and 10 h15 �C/light off; the relative humidity ranged between 60 and 70%.Light was provided by 16 metal-halide lamps (HQI e 400 Watt,Osram); the photosynthetic photon flux density ranged from450 mmol photon m�2 s�1 at the pot level to 1600 mmol photonm�2 s�1 at the ceiling. Nutrients were supplied as liquid complexsolution (Hoagland) everyweek, according to the growth rate of theplants. At maturity, leaves were forced to fall by reducing the lightperiod and collected after falling.

In October 2004, two PVC collars (10 cm high, 20 cm diameter)were placed in the centre of P. nigra stands (planting density 1�1m),within each of the 2 sectors of the 6 plots. The collars were insertedinto the soil to 2 cm depth and all the native litter inside the collarswas removed. Ten grams of dried 13C-labelled litterwas placed on topof the mineral soil as a treatment. No litter was applied in equivalentcontrols. Within each large PVC collar, one small PVC collar (10 cmhigh,11 cmdiameter) was inserted into the soil to 2 cm depth for soilCO2 effluxmeasurements. The collars were covered by a coarse high-density PVC net (mesh size: 1 cm � 1 cm) in order to prevent freshlitter from falling into the collars.

2.2. Soil CO2 efflux measurements

The modified procedure of Subke et al. (2004) was used formeasuring soil CO2 efflux and capturing the CO2 for subsequentisotopic analysis was adopted. Soil CO2 efflux measurements werecarried out on a monthly basis over 11 months. A portable infraredgas analyser (IRGA) EGM-4 (PP Systems, Hertfordshire, UK) wasconnected through a closed type cuvette SRC1 (PP Systems, Hert-fordshire, UK) to the internal collar of each plot. The rate of soil CO2

efflux was obtained by the software of the PP system, which fitteda linear equation to the increase of CO2 concentration in the cuvetterecorded every 120 s. The measurement was repeated 3 times foreach collar, with the chamber being displaced and repositioned onthe collar at each repetition. The mean value of the 3 repeatedmeasurements was taken as a reading for each collar. The soiltemperature at 10 cm depth was measured for each plot at the time

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e1016 1011

of efflux measurement. Efflux measurements from all collars wereconducted on the same day at each sampling occasion.

2.3. Sampling of respired CO2 and d13C measurementof the soil CO2 efflux

The sampling of the respired CO2 for 13C analyses was carriedout on the same day of soil CO2 efflux measurement. To accumulatethe respired CO2 for sampling, a PVC lid was placed on each collar,with an internal o-ring made of teflon and an external teflon tubingconnection (6 mm internal diameter). A 2 L previously evacuatedflask was connected to the lid through the external tubing. Thechamber was left for 20 min to allow an increase head space CO2 bydiffusion from the soil. The initiation of this enrichment period wasat least 30 min after the flux measurement had been completed ona collar, to ensure that no disturbance of the concentration gradientpersisted. After the 20 min enrichment period, the flask was filledwith air coming from the chamber. On each sampling day, oneadditional air samplewas taken for each of the 6 experimental plotsfor correction of the d13C values of chamber air samples with regardto atmospheric d13C values (Equation (2)).

2.4. Litter collection and soil sampling

In September 2005, all recognizable litter residues on the soilsurface within each collar were collected. 20 � 20 cm soil sampleswere taken from beneath all PVC collars and divided into 4 depthhorizons: 0e2, 2e5, 5e10,10e20 cmgiving 96 samples (4 depths� 2collars � 2 sectors � 6 plots). The collected litter residues and soilsamples were oven dried at 70 �C to constant weight and ground topass a 500 mm mesh for analysis.

2.5. Chemical and isotopic analyses

Soil samples were treated with HCl to avoid any contaminationby inorganic C, after Harris et al. (2001). C and N concentrationswere measured using an elemental analyser (Thermofinnigan EA1112), and bulk d13C values were measured using an IRMS (Ther-mofinnigan Deltaplus) coupled to an elemental analyser (Thermo-finnigan EA 1112). The d13C values (&) of CO2 in air samples wasmeasured by cryogenic extraction of the CO2 and subsequentanalyses with the IRMS (Bertolini et al., 2005).

2.6. PLFA analyses

Bulk d13C analysis of the four soil layers revealed that only the0e2 cm layer was significantly 13C-enriched (Fig. 2), therefore,only this horizon was used for PLFA analysis. Ten grams of freezedried, ground soil were extracted using a modified Bligh-Dyermonophasic solvent system (Crossman et al., 2004). Briefly,

Table 1Groups of micro-organisms associated with production of characteristic types of phospholet al., 2001; Ohtonen et al., 1999; Olsson, 1999; Sundh et al., 1997; Zak et al., 1996; Zelle

Abbreviations Types of FAs Ex

EL-SATFAs (Ester linkedSaturated Fatty Acids)

Branched i14Cyclopropyl cyMethyl branchedon 10th C atom

10

Long, straight chain 20EL-MUFAs (Ester Linked

Monounsaturated Fatty Acids)Straight chain 16u8 1616:1u5 1618:1u9 18

EL-PUFAs (Ester LinkedPolyunsaturated Fatty Acids)

all PUFA all18:2u6,9 18

Bligh-Dyer solvent containing water buffer (0.05 M KH2PO4; pH7.2)-chloroform-methanol in a ratio of 4:5:10 (v/v/v) was addedto each soil sample. The mixture was ultrasonicated (15 min) andcentrifuged (3000 rpm, 5 min). The supernatant was decanted,and the residue re-extracted three times. The extracts werecombined and evaporated under N2 to yield the total lipidextract, which was separated into three fractions based onpolarity using a 3 ml STRATA aminopropyl-bonded silica solidphase extraction cartridge (Phenomenex). Fractions were elutedsequentially with 2:1 (v/v) dichloromethane-isopropanol (8 ml;yielding neutrals), 2% acetic acid in diethyl ether (12 ml; freefatty acids), and methanol (8 ml; polar fraction including phos-pholipids). The PLFA components of the polar fraction werereleased by saponification (1 ml 0.5 M methanolic NaOH, 1 h,80 �C), and an internal standard was added (20 ml 0.1 mg ml�1 C19n-alkane solution in hexane). The PLFAs were then methylatedusing a 14% (w/v) BF3-methanol solution (100 ml; Aldrich; 1 h,80 �C) and extracted with chloroform (3 portions of 2 ml). Thefatty acid methyl esters (FAMEs) were dissolved in hexane foranalysis by GC, GCeMS, and GC-C-IRMS.

FAMES were analysed on a Hewlett Packard 5890 Series II gaschromatograph. Sample introduction was via on-column injector,and the detector was a flame ionisation detector. FAME separationwas achieved using a Varian Factor Four VF23MS column(60m� 0.32mm [i.d.]; 0.15-mm film thickness). The carrier gas wasH2 (10 p.s.i. head pressure), and the oven temperature programmedfrom 50 �C (1 min isothermal) to 150 �C at 15 �C min�1, from 150 �Cto 240 �C at 4 �C min�1, and held at 240 �C for 20 min isothermal.Quantification of components present in the fatty acid GC profileswas performed relative to the internal standard.

FAMES were identified using a Thermofinnigan Trace GCeMSsystem. The column and temperature program details were thesame as those described above except He was used as the carriergas. The interface was set to the maximum oven temperature, theion source was set to 30 �C below this and the quadropole MSscanned the range of m/z 50 to 650 at 1.7 scans s�1. Data wereacquired and processed using Excalibur data system Version 1.2.Biomarker PLFAwere interpreted according to a range of previouslypublished references (Table 1).

The d13C values of FAMES were determined using a Varian 3400GC attached to a Finnigan MAT Delta S IRMS (electron ionisation,100 eV, 1 mA electron energy, 3 Faraday cup collectors m/z 44, 45and 46, CuO/Pt Finnigan MAT Mark I combustion interface main-tained at 850 �C). Water removal was via a Nafion membrane. GCconditions were the same as those described above except that Hewas the carrier gas. Samples were calibrated against reference CO2of known isotopic composition, introduced directly into the sourcethree times at the beginning and end of every run. Each sample wasrun in duplicate to ensure reliable mean d13C values. FAME refer-ence standards were run every four samples.

ipid fatty acids (derived from: Frostegard et al., 1997; Frostegard et al., 1993; Graystons, 1999, 1997).

amples Biomarkers for

:0, i15:0, a15:0, i16:0, i17:0, i18:0 mainly gram-positive17:0, cy19:0 mainly gram-negativeMe16:0, 10Me17:0, 10Me18:0 actinomycetes

:0, 21:0, 22:0, 23:0, 24:0, 25:0, 26:0 higher plants, mosses:1u9, 16:1u7, 18:1u7 mainly gram-negative:1u8, 18:1u8 methane-oxidizing bacteria:1u5 arbuscular mycorrhyzal fungi:1u9 mainly in fungiPUFA do not occur in bacteria:2u6,9 mainly in fungi

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e10161012

2.7. Calculations and statistical analysis

The experiment was conductedwithin the experimental plots ofthe EUROFACE project (Scarascia Mugnozza et al., 2000), however itwas carried out after the CO2 fumigation had been stopped. Thus,for the purpose of this study, and because at the end of the incu-bation experiment no significant difference among either sectors orplots was found (P < 0.01), the samples were treated as 12 inde-pendent replicates (12 with litter treatments and 12 control soils).A Student’s t-test was used to explore differences between values atthe beginning and at the end of the experiment. In all casesconfidence limits in excess of 95% were considered significant.

2.7.1. SOM-CThe fraction of litter-derived C in the different soil layers ðCl

s=CsÞwas obtained by applying a two source mixing model of thedifference in d values between the with litter soil (ds) and thecontrol soil (dc) at the end of the experiment, according to:

Cls

Cs¼ ds � dc

dl � dc(1)

where dl is the d13C value of the litter sample. It was assumed thatthe d13C values of the control soils were equivalent to the d13Cvalues of the initial soil C prior to incubationwith 13C-labelled litter,and that the d13C values of the litter-derived C incorporated intoSOM was equivalent to the d13C values of the bulk litter.

2.7.2. CO2eCThe d13C values of the CO2 from each ring were corrected using

the d13CvsPDB of the atmospheric sample taken at the same ring, oneach sampling date, according to:

d13Cr � ½CO2�r ¼ d13Cc � ½CO2�c�d13Ca � ½CO2�a (2)

where the subscript “r”, “c” and “a” refer to the CO2 “respired”, thetotal CO2 measured in the “chamber” and the contribution from the“atmosphere”, respectively.

The CO2 concentrations of air samples were determined bycomparing the m/z 44 signal at the Faraday cup of the IRMS for thecryogenized CO2 present in samples to the signal obtained froma standard air cylinder with known CO2 concentration and isotopiccomposition. A constant cryogenization efficiency was assumedand, was not significantly different from 100% (Bertolini et al.,2005). Different known amounts of CO2 from the standardcylinder were cryogenized and analysed to create a calibrationcurve: mol of CO2 vs signal at m/z 44. The concentration of CO2 inunknown samples was obtained by interpolation with a linear fit.

With respect to the CO2 respired in the SOM, roots and therooting environment were considered as one isotopic pool (indi-cated by the suffix “s” for “soil”), the fraction of CO2 produced by thelitter, over the total CO2 respired by the system, was, therefore,estimated according to:

COl2

COt2¼ dCOt

2 � dCOs2

dCOl2 � dCOs

2

(3)

where COl2=CO

t2 is the fraction of CO2 respired by the litter over the

total CO2 respired by the soil and the litter; dCOt2 represents the

isotopic composition of sampled air CO2; dCOs2 is the isotopic

composition of the CO2 respired by the control soil; dCOl2 is the

isotopic composition of the CO2 respired from the litter. The mainassumption of this is that no isotopic fractionationwas associated tothe respiration process ðdCOl

2 ¼ dl; dCOs2 ¼ dcÞ. To validate this

assumption, the dCOl2 was measured by incubating litter samples of

ca. 3 g in 1 L jars, in three replicates, and cryogenically trapping therespired CO2, following themethod described in Rubino et al. (2007).

The CO2 flux measurements were integrated over time to obtainthe total quantity of CO2 respired by the samples during the entireincubation period. An exponential relationship ðR ¼ aektÞ wasapplied between CO2 fluxes (R) and soil temperature (T) and thebest estimates for a and k were found on the basis of ourmeasurements. The estimated parameters were used to derive thehalf hourly soil CO2 efflux for the entire experiment from themeasured daily soil temperature (data not shown). Cumulative CO2respired by litter samples for the entire duration of the litterdecomposition experiment was obtained by numerical integrationand partitioning according to Equation (3).

2.7.3. PLFA-Cd13C values obtained for the methylated fatty acids were cor-

rected for the addition of the derivative C using the mass balanceequation shown in Equation (4). The d13C value for the BF3-MeOH(d13C ¼ �34.1&) was determined by bulk CF-IRMS:

ncdd13Ccd ¼ ncd

13Cc þ ndd13Cd (4)

where “n” is the “number” of C atoms, “c” is the “underivatisedcompound”, “d” is the “derivatising agent”, and “cd” is the “deriva-tised compound” of interest.

Fatty acids structure is represented by two numbers separatedby a colon (see Table 1). The number before the colon indicates theC chain length and the number after corresponds to the number ofdouble bonds. The position of the first double bond is counted fromthe methyl end of the molecule and prefixed by the symbol u.Methyl branching on the first (iso) or second (anteiso) C is desig-nated by i or a prefixes, respectively.

The fraction of litter-derived C in different PLFAs ðCls=CsÞ was

determined using the difference in d values betweenwith litter-soilðdsÞ and control soil ðdcÞ at the end of the experiment, according to:

Cls

Cs¼ ds � dc

dl � dc(5)

where the symbols have the same meaning than the symbols inEquation (1), but are measured for each specific PLFA.

The absolute quantity of litter-derived C for each PLFA fromlitter-treated soil was obtained by multiplying the percentage oflitter-derived C in each PLFA by the amount of total C in the cor-responding PLFA. We referred this quantity to the C lost by the leaflitter samples in order to quantify how much of the litter C wasincorporated into each group of PLFA.

3. Results

The variations, as mean of the 12 replicates, in terms of littermass loss, C concentration and d13C values between the beginningand the end of the experiment are reported in Table 2.

Eighty per cent of initial mass was lost over the duration of theexperiment. The N content of the litter was significantly increased(P < 0.01) with respect to the initial value. This increase wasreflected in a decrease of C/N ratios. Additionally, a significant(P< 0.01) depletion in 13C was measured for all litter samples at theend of the experiment.

3.1. CO2 efflux and its components

Monthly respiration rates for litter-treated and control soils areshown in Fig. 1a. A significant difference was found between dates(P < 0.01) indicating a seasonal trend, with good linear correlation

Table 2Mass [g], C and N concentrations [%], C/N ratio and d13C [&] values of leaf litters atthe beginning and at the end of the experiment. Mass (Dm) and C (DC) loss [%] overthe total initial mass and C amount, respectively, and amount [g] and fractions [%] ofC lost by litter samples as C input to the belowground SOM (SOM-C) and as CO2

(CO2eC) over the total litter C loss. Numbers in brackets are standard errors overn ¼ 12 replicates.

Initial Final D

m [g] 9.46 (0.02) 1.9 (0.3)C [%] 43.4 (0.3) 43 (1)N [%] 0.25 (0.03) 2.2 (0.1)C/N 173 (22) 19 (1)d13C [&] 166.3 (0.7) 125 (6)

m [%] 80 (3)C [g] 3.3 (0.1)

SOM-C [g] 2.2 (0.4)CO2eC [g] 1.0 (0.1)SOM-C [%] 67 (12)CO2eC [%] 30 (3)

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e1016 1013

between the mean respiration rates for each month, except July,and the mean soil temperature: R ¼ 0.843 (P < 0.05). It is worthnoting that the soil at the site was not limited by moisture(Scarascia Mugnozza et al., 2000).

The mean d13C value of the CO2 respired by litter in the labo-ratory incubation was 169 � 7& (mean � standard error, n ¼ 3)which was not significantly different (P < 0.01) from the d13C valueof the bulk litter. There was no significant difference (P < 0.05)between the d13C values of CO2 respired by the control soils duringthe entire study. For this reason, they are schematically representedby three parallel lines (mean value � standard error overall

0

10

20

-30

-20

-10

0

10

20

30 frettil

]%[

31O

C-C

2]

‰[B

DPsv

Control soil Control soil ± std.err.

Litter-treated flitter [%]

0

5

10

15

20

25

Tlios

]C°[

Tmean

0

3

6

OC

2[ xulffe

OClo

m μ δ

2s

1-m

2-]

Litter-treated

a

b

Control soil

Fig. 1. a) monthly respiration rate [mmol CO2 s�1 m�2] for litter-treated (white bars)and control soils (black bars). Black circles represent mean soil temperature on eachsampling day; b) d13CeCO2 [&]vsPDB of litter-treated (white triangles) and control soil(black line) and fraction (black triangles) of litter-derived C in the CO2 respired bylitter-treated soil. Error bars are standard errors over n ¼ 12 replicates.

measured values) in Fig. 1b. At the beginning of the experiment, thed13C values of CO2 respired by litter-treated soils (Fig. 1b) weremore positive than the corresponding values for control soils,indicating a significant contribution of litter to the respirationefflux. The d13C values became more negative with time and werenot significantly different from those of control soils in June, whenlitter had already visibly decomposed. As a consequence, the littercontributions to the total CO2 efflux (Fig. 1b) had a similardecreasing trend. In conclusion, about one-third (30 � 3%) of thetotal litter C was lost as CO2.

3.2. C input from aboveground leaf litter to belowground SOM

The d13C values of litter-treated soils were progressively13C-depleted with depth (Fig. 2). Conversely, the control soilshowed a more uniform d13C distribution down the profile andwith more 13C-enriched values nearer the surface. Only the top twosoil horizons (0e2 and 2e5 cm) of the litter-treated soils weresignificantly enriched in 13C compared to the control (P< 0.01). Thefraction of litter-derived C was 5% of the total C in the soil layer(Fig. 2). Overall, it was calculated that around two-thirds (67 � 13)of the C in the initial litter had been incorporated into the SOM after11 months.

3.3. PLFA abundance and d13C values

Fig. 3 shows the distribution of PLFAs extracted from one of thereplicates of the litter-treated soil at the end of the experiment.Apart from16:0, themost abundant PLFAswere branched saturatedfatty acids (SATFA) i15:0, a15:0, i16:0, i17:0, and a17:0, character-istic of Gram-positive (Gþve) bacteria, compared to other microbialgroups Gram-negative (G�ve) bacteria (16:1u7/9, 18:1u7/9,cy17:0/19:0), actinomycetes (10Me16:0/18:0) and fungi (18:2u6,9).

The distribution of PLFA extracted from the litter-treated soiland control soil was very similar. However, there were significantdifferences in PLFA concentration (mg C g�1soil) between microbialgroups (P < 0.01; Fig. 4a): Gþve bacteria were the most abundantgroup of micro-organisms identified.

A significant enrichment in 13C was determined in all PLFAs fromthe litter-treated plots compared to the controls (Fig. 4b), revealingsustained incorporation of litter-derived C in the SMB after 11months. The d13C values of PLFAs were not significantly differentamong groups of micro-organisms, with 7.8 � 1.6% of total PLFA-Cbeing litter-derived. As the most abundant group identified, Gþve

-30 -20 -10 0 10

depth [cm]

δ13C [‰]vsPDB f [%]

Control soil

Litter-treated

0-2

2-5

5-10

10-20

Fig. 2. d13CeCO2 [&]vsPDB (left side) of litter-treated soil (white squares) and controlsoils (black squares) and fraction (%, right side) of litter-derived C in soil for eachsampled soil layer (0e2, 2e5, 5e10, 10e20 cm). Error bars are standard errors over 12replicates.

Fig. 3. Partial TIC chromatogram of methyl esters of PLFAs extracted from litter-treatedsoil. Key: 1 ¼ internal standard (n-nonadecane); 2. i14:0; 3. 14:0; 4. i15:0; 5. a15:0;6. 15:0; 7. i16:0; 8. 16:0; 9. cy17:0; 10. 10Me16:0/16:1u9; 11. 16:1u7; 12. i17:0;13. a17:0; 14. 17:0; 15. br18:0, 16. i18:0; 17. 18:0; 18. cy19:0; 19. 10Me18:0; 20. 18:1u9;21. 18:1u7; 22. C18:2u6,9; 23. i20:0; 24. a20:0; 25. 20:0.

0

1

2

3

g Cg

1-lios

Litter-treated

Control soil

0

20

40

60

Gram + Gram - Actinomycetes Fungi

μ μ

g Cg

1-C-deviredrettil

-35

-25

-15

-5

δ 31C

BDPsv

]‰[

a

b

c

Fig. 4. a) PLFA concentrations [mgPLFA-C g�1soil] for litter-treated (white bars) and control

soil (black bars); b) PLFA d13CeCO2 [&]vsPDB for litter-treated (white squares) and controlsoil (black squares); c) concentration of litter derived PLFA-C [mgC g�1

C lost from litter] overthe total C lost by the litter. Error bars are standard errors over 12 replicates.

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e10161014

bacteria incorporated a higher concentration of litter-derived Cthan G�ve bacteria, actinomycetes and fungi (Fig. 4c).

4. Discussion

4.1. Partitioning of C loss

The C loss from the litter, determined as components of SOMand CO2, accounted for 97 � 13% of the total C in the initial litter(Table 2). The large standard deviation (13%) was expected ina highly variable system, reflecting the complex nature of theinteractions between litter, roots, microbes and nutrients supply, asalready underlined by Bending (2003) and Ross et al. (2002).

The ratio of C lost as CO2 to C input to SOM was around 1:2.Overall we identified a larger mass loss than previously deter-mined. Cotrufo et al. (2005) performed a litter bag decompositionstudy at the same site, and concluded that Populus sp. litter only lostbetween 15 and 18% of the original mass during 8 months of fieldincubation. Litter bag studies prevent fragmentation and subse-quent transport, and consequently mass loss is largely a result ofmicrobial degradation of litter within the litter itself, therebyunderestimating decay rates. Indeed, our study shows that thisprocess was not the major factor contributing to C input to SOMduring litter decay, underlining the need to account for litter frag-mentation in aboveground litter decomposition studies (Kheirallah,1990). Additionally, litter enters the soil in the form of undecom-posed litter fragments (i.e. light fraction) which can initiate SOMaggregation and thus play a pivotal role in the C stabilizationmechanisms (Del Galdo et al., 2003; Six and Jastrow, 2002).Furthermore, fragmented pieces of litter within soil are likely todecompose at a rate that is different from the decomposition rate ofeither surface litter or humified SOM (Adair et al., 2008). This showsthe requirement to view SOM as a continuum of different states ofdecomposition which could have important implications for con-straining terrestrial C pools.

In addition to the free movement of fragmented plant matter,the incubation of plant litter in direct contact with the soil allowsmore natural interaction with soil fauna. The effect of incubationwithin collars on the ecology of decomposing macrofauna mayhave favoured those entering from the soil and also reducedpotential predation. Nonetheless, we assumed that any effectassociated with the experimental systems had a smaller influenceon the decomposition process than the underestimation caused bythe litterbags methodology, as quantified by Bradford et al. (2002),Smith and Bradford (2003) and St. John (1980).

4.2. N and C dynamics during litter decomposition

Nitrogen concentration had increased significantly in the litterby the end of the experiment (Table 2), with an average net increaseof 1.9 � 0.7 mg N g�1 litter probably occurring by uptake andimmobilization of external N in leaf litter during decomposition(Qinglin et al., 2009). Atmospheric deposition in the region of thestudy site was not sufficient to explain the increase (source: http://www.bgc.mpg.de/public/carboeur/sites/rocca.html), and no signif-icant difference was found between N-fertilised and control plots,excluding an influence of the N fertilization. Therefore, the increaseis suspected to be via the incorporation of N within the microbialdecomposer community within the litter, particularly fungi (Freyet al., 2003).

Co-transport of organic N by microbes from soil which isdepleted in 13C compared to litter had previously been reported by(Aikio et al., 2000; Gregorich et al., 1991). However, variation in thed13C value of litter during decomposition might also be a conse-quence of isotopic fractionation during the respiration process

M. Rubino et al. / Soil Biology & Biochemistry 42 (2010) 1009e1016 1015

operated by micro-organisms, causing the preferential release of13C (Connin et al., 2001; Fernandez et al., 2003; Schweizer et al.,1999). Selective degradation of different compounds with a rangeof d13C values over time may also explain the variation in d13Cvalues, as compounds considered to be more recalcitrant arereported to be relatively 13C-depleted, i.e. lignin and lipids(Fernandez et al., 2003; Schweizer et al., 1999). However, deter-mining the processes acting to alter d13C values of litter C requiresmore detailed analyses which is beyond the scope of his paper.

4.3. Effect of litter application on the SMB

The population profile of the SMB, determined using GC anal-yses of PLFA, appeared similar to that prior the application of13C-labelled P. nigra, which replaced native P. nigra litter. Evidenceof litter C incorporation could, therefore, only be detected through13C-PLFA analysis by GC-C-IRMS. PLFA concentration andcompound specific d13C values were used to calculate the % w/w oflitter C incorporated into the different microbial groups, i.e.: Gþveand G�ve bacteria, actinomycetes and fungi (Equation (5)). Allgroups contained ca. 8% PLFA-C assimilated from 13C-labelled litter(Fig. 4b) suggesting that all groups were able to incorporate litter Cat a rate proportional to their contribution to the SMB. Significantly,Gþve bacteria were the dominant group of micro-organisms in thesoil, and contained w75% w/w litter-derived C assimilated bythe SMB after one year (Fig. 4c). Gþve bacteria are reported to bemore able to degrade complex substrates, but have slower growthrates than G�ve bacteria (Kuzyakov et al., 2000), so their prolifer-ation may be an adaptation to more biochemically complex,partially decomposed litter input. Therefore, it might be expectedthat the d13C values of Gþve bacteria would be different comparedto less active microbial groups who rely on more labile substratesavailable early in decomposition. However, Fig. 4 shows a similarmean d13C value of around �12.5 � 3.1& for Gþve, G�ve, actino-mycetes and fungal PLFA. This suggests that either, (i) all micro-organisms were incorporating a similar component of litter C at thesame rate, or, (ii) after 11 months, the system was at steady statesuch that incorporated litter C was being recycled within the SMBgiving rise to homogenous d13C values between PLFAs.

Though this study was limited to 2 dates at the beginning andend of the experiment, we have shown that the quantification ofthe amount of C incorporated by different groups of soil micro-organisms is a useful approach. This study can be developed bothon a temporal basis to investigate microbial succession active in thelitter decomposition at higher time resolution and with the regardsto the input of various soil C compounds both from the litter(Derrien et al., 2003; Dignac et al., 2005; Gleixner et al., 2001) andas metabolites from the microbial biomass itself (Rubino et al.,2007).

5. Conclusion

In this experiment, we determined that two-thirds of the litter-derivedC input to themineral soil probablyentered the soil in solubleor fragmented form, whilst the fraction of CeCO2 loss was one-thirdof total C lost from litter for all treatments. Sustained incorporationand turnover of litter C by the SMB was determined using 13C-PLFAanalysis after 11 months, revealing a mechanism for continuedmineralisation and turnover of litter C in short-rotation coppiceplantations. Unlike most previous studies using litterbags, we haveshown that litter C incorporation by the microbial biomass withinthe litter may not represent the major flux during decomposition.From a field-based, methodological standpoint, these results suggestthat the use of litterbags should be avoided, since bags significantlyalter the flow of C from decomposing litter to the SOM and, thus,

prevent anaccurate estimate of surface littermass loss anddecay ratein natural systems.

Future measurements should clarify the proportion of litter-derived C incorporated into the range of SOM fractions, e.g.: lightfraction or particulate organic matter, and its biomolecularcomposition, allowing a more holistic understanding of the litter-soil biogeochemical continuum.

Acknowledgments

The work was supported by the EUROFACE project (EVR1-CT2002-40027) and co-financed by Legge 5, Regione Campania,Italy. We want to thank Jens Arne Subke for his technical sugges-tions about CO2 efflux measurements and Francesco Pascale for hisgreat help during field work at sample collection. The use of theNERC Life Sciences Mass Spectrometry facilities, particularlythe technical support from Drs Ian Bull and Rob Berstan, at theUniversity of Bristol is gratefully acknowledged.

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