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Soil Biology & Biochemistry 41 (2009) 144–153

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Soil Biology & Biochemistry

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

Microbial community composition and rhizodeposit-carbon assimilationin differently managed temperate grassland soils

Karolien Denef a, Dries Roobroeck a, Mihiri C.W. Manimel Wadu b, Peter Lootens c, Pascal Boeckx a,*

a Laboratory of Applied Physical Chemistry, Ghent University, 9000 Gent, Belgiumb Interuniversity Graduate Degree Program in Physical Land Resources, Ghent University, Belgiumc Institute for Agricultural and Fisheries Research (ILVO), 9820 Merelbeke, Belgium

a r t i c l e i n f o

Article history:Received 7 February 2008Received in revised form30 September 2008Accepted 6 October 2008Available online 31 October 2008

Keywords:GrasslandPulse-labelingMicrobial community structurePLFA13CMowingFertilization

* Corresponding author.E-mail address: pascal.boeckx@ugent.be (P. Boeck

0038-0717/$ – see front matter � 2008 Elsevier Ltd.doi:10.1016/j.soilbio.2008.10.008

a b s t r a c t

Rhizodeposit-carbon provides a major energy source for microbial growth in the rhizosphere of grass-land soils. However, little is known about the microbial communities that mediate the rhizospherecarbon dynamics, especially how their activity is influenced by changes in soil management. Wecombined a 13CO2 pulse-labeling experiment with phospholipid fatty acid (PLFA) analysis in differentlymanaged Belgian grasslands to identify the active rhizodeposit-C assimilating microbial communities inthese grasslands and to evaluate their response to management practices. Experimental treatmentsconsisted of three mineral N fertilization levels (0, 225 and 450 kg N ha�1 y�1) and two mowingfrequencies (3 and 5 times y�1). Phospholipid fatty acids were extracted from surface (0–5 cm) bulk (BU)and root-adhering (RA) soil samples prior to and 24 h after pulse-labeling and were analyzed by gaschromatography–combustion-isotope ratio mass spectrometry (GC–c-IRMS). Soil habitats significantlydiffered in microbial community structure (as revealed by multivariate analysis of mol% biomarkerPLFAs) as well as in gram-positive bacterial rhizodeposit-C uptake (as revealed by greater 13C-PLFAenrichment following pulse-labeling in RA compared to BU soil in the 450N/5M treatment). Mowingfrequency did not significantly alter the relative abundance (mol%) or activity (13C enrichment) ofmicrobial communities. In the non-fertilized treatment, the greatest 13C enrichment was seen in allfungal biomarker PLFAs (C16:1u5, C18:1u9, C18:2u6,9 and C18:3u3,6,9), which demonstrates a prom-inent contribution of fungi in the processing of new photosynthate-C in non-fertilized grassland soils. Inall treatments, the lowest 13C enrichment was found in gram-positive bacterial and actinomycetesbiomarker PLFAs. Fungal biomarker PLFAs had significantly lower 13C enrichment in the fertilizedcompared to non-fertilized treatments in BU soil (C16:1u5, C18:1u9) as well as RA soil (all fungalbiomarkers). While these observations clearly indicated a negative effect of N fertilization on fungalassimilation of plant-derived C, the effect of N fertilization on fungal abundance could only be detectedfor the arbuscular mycorrhizal fungal (AMF) PLFA (C16:1u5). On the other hand, increases in the relativeabundance of gram-positive bacterial PLFAs with N fertilization were found without concomitantincreases in 13C enrichment following pulse-labeling. We conclude that in situ 13C pulse-labeling of PLFAsis an effective tool to detect functional changes of those microbial communities that are dominantlyinvolved in the immediate processing of new rhizosphere-C.

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1. Introduction

Soil microorganisms are crucial for the maintenance of soilfertility and sustainability of grassland ecosystems due to theirimportance in regulating the decomposition of organic matter andnutrient cycling. Their habitat in grassland soils can be described asa dynamic and heterogeneous environment characterized by

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numerous abiotic and biotic processes which can drastically changeunder changes in land-use, management or environmental condi-tions. This could cause shifts in microbial growth and activity aswell as in microbial community composition. The latter may havea considerable feedback on key biogeochemical processes in soils.For instance, it has been postulated that increases in the abundanceof fungi relative to bacteria in the soil microbial community areassociated with reduced rates of nutrient cycling (e.g. N minerali-zation) and increased retention of C and N in the soil (Bardgett,2005; de Vries et al., 2006; Hogberg et al., 2007). Shifts in microbialcommunity composition in response to grassland management

Table 1Mineral N application (kg N ha�1) and mowing events during 2006. Time of pulse-labeling is indicated.

Date 0N/3M 225N/3M 225N/5M 450N/3M 450N/5M

20 April 2006 75N 50N 80N 100N2 May 2006 70N4 May 2006 Mowing Mowing9 May 2006 50N 100N16 May 2006 Mowing Mowing Mowing24 May 2006 75N 80N5 June 2006 Mowing Mowing12 June 2006 45N 70N 90N24 July 2006 Mowing Mowing Mowing Mowing Mowing28 July 2006 75N 40N 80N 80N23–28 August 2006 Pulse-labeling4 September 2006 Mowing Mowing8 September 2006 40N 70N 80N9 October 2006 Mowing Mowing Mowing Mowing Mowing

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153 145

have been demonstrated by many researchers. Generally, a shift insoil microbial community structure has been found from onefavoring fungi to one favoring bacteria as grassland managementintensity increases (e.g. Donnison et al., 2000; Grayston et al.,2001). Fungi in particular appear to be very sensitive to manage-ment practices (Zeller et al., 2000) such as mineral fertilizer addi-tion (Bardgett et al., 1996), manure addition (Bittman et al., 2005)and grazing (Bardgett et al., 2001). Mowing has been found tostimulate mycorrhizal fungal growth (Antonsen and Olsson, 2005),most likely as a result of increased root respiration and carbonexudation (Mawdsley and Bardgett, 1997).

Soil microorganisms heavily depend on organic C supply fortheir growth. Plants provide carbon to rhizosphere microorganismspartly in the form of rhizodeposits which comprise exudates (smallmolecules, such as organic acids, amino acids and sugars), secre-tions (such as enzymes), lysates from dead cells, and mucilage(Singh et al., 2004). Despite being a small proportion of the total Cinputs from vegetation to soil, rhizodeposits are believed to benefitthe microbial community in the rhizosphere (Jones et al., 2004;Singh et al., 2004). However, there exists less evidence about whichorganisms benefit most from new rhizodeposit-C inputs, and howtheir activity is affected by management or changes in environ-mental conditions. For this, in situ studies are needed that trace theflow of C from plants to active C assimilating microorganisms. Onedirect approach to determine the C flow through the plant–soil-microbial system involves the exposure of photosynthesizingplants, either in the field or laboratory, to 13C-labeled CO2 (Ostleet al., 2000; Leake et al., 2006). The isotopic label can then be tracedin microbial biomarker molecules to identify the microorganismsthat directly or indirectly metabolize the newly deposited plant-derived C. A number of recent studies have used this 13CO2 pulse-labeling technique to label microbial phospholipid fatty acids(PLFAs) in order to assess the activity of rhizosphere microbialcommunities (Butler et al., 2003; Treonis et al., 2004; Denef et al.,2007; Lu et al., 2007). These studies have demonstrated the pres-ence of an active microbial community in the rhizosphere of grassesand rice; different contribution to the assimilation of rhizodeposit-C by different microbial communities; and temporal and spatialvariations in C assimilation by microbial communities as well asvariations with changes in grassland management (e.g. liming,Treonis et al., 2004) or environment (e.g. elevated CO2, Denef et al.,2007).

The aim of this particular study was to investigate the impact ofinorganic N fertilizer addition and mowing frequency on theabundance and activity of microbial communities in temperategrassland soils. The combined effect of these two managementpractices was investigated since mowing frequency usuallyincreases with increased fertilizer addition in managed grasslandsdue to higher productivity. Our study combined an in situ 13CO2

pulse-labeling approach with 13C-PLFA analysis to assess whichmembers of the microbial community actively assimilate new rhi-zodeposit-C and how their activity is affected by N fertilizer addi-tion and mowing frequency. 13C-PLFA results were comparedbetween two soil habitats to detect any spatial variation of activemicroorganisms associated with the rhizosphere carbon flow. Theeffects of grassland management on microbial rhizodeposit-Cuptake were expected to be greatest near the roots where therhizodeposit-C utilizing microbial community was expected to bemost active.

2. Materials and methods

2.1. Experimental site and treatments

The study sites were located at the experimental research areaof the Institute for Agricultural and Fisheries Research (ILVO) near

Merelbeke (Belgium) (50.97330�N; 3.78119�E). Soils are classifiedas sandy loam (Belgian soil classification). This area was used ascropland for decades prior to establishment of grassland. In spring2001, the cropland was converted to typical grassland for agricul-tural use with an established vegetation mixture composed ofLolium perenne ‘‘Ritz’’ (35%), L. perenne ‘‘Pandora’’ (35%), Phleumpratense ‘‘Erecta’’ (15%), Festuca pratensis ‘‘Merifest’’ (10%) and Poapratensis ‘‘Balin’’ (5%). Five treatments were selected, each con-sisting of a specific combination of mineral N fertilization level andmowing frequency: 0 kg N ha�1 y�1 with 3 mows per year (0N/3M);225 kg N ha�1 y�1 with 3 mows (225N/3M) and 5 mows (225N/5M)per year; 450 kg N ha�1 y�1 with 3 mows (450N/3M) and 5 mows(450N/5M) per year. In all treatments, hay was removed aftermowing. The time schedule for N fertilizer addition and mowingduring 2006, i.e. the year in which the pulse-labeling experimentwas performed, is given in Table 1. Mineral N was applied as part ofa NPK (15-7-22) fertilizer mixture with 6.2% N as nitrate and 8.8% Nas ammonia. The experiment had a split-plot design with fourblocks. Main plots consisted of N fertilizer treatment. Subplots(6.3 m� 1.2 m) consisted of mowing frequency treatment. Mainplots and subplots were not randomized within each block.

2.2. 13CO2 pulse-labeling experiment

Four replicate 13CO2 pulse-labeling events were performedbetween the 23rd and 28th of August 2006. Per pulse-labelingevent, a 625 cm2 area in each of the 5 selected treatment plots perblock was labeled by injecting 13C-labeled CO2 (99 atom%; LansMedical, The Netherlands) into closed top plexiglass chambers(25 cm� 25 cm� 25 cm). One pulse-labeling event was performedper replicate block (labeling one replicate of each of the 5 treat-ments), and thus repeated 4 times (4 different pulse-labeling days).One day prior to each pulse-labeling event, steel collars wereinserted into the soil to a depth of about 10 cm. At the advent ofa pulse-labeling event, the chambers were positioned on top ofthese collars containing a thin layer of water to seal the chambers.The CO2 concentration was monitored with an EGM-4 (PP systems,Massachusetts, US). When the initial CO2 level in the chamberdropped below 250 ppm, 13C-labeled CO2 was injected into thechambers to obtain a theoretical increase of 1000 ppm in CO2

concentration. This was repeated throughout a 6-h period, based onthe initial CO2 uptake rate in each chamber. Since the CO2

concentration decreased at different rates in the different treat-ment plots, not all chambers received the same amount of 13C.Notwithstanding, a constant amount of 13C was added to all repli-cate pulse-labeling chambers of a particular treatment. Thechambers were only removed from the grass areas 24 h after the

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153146

first injection of 13CO2. This was done to ensure maximum incor-poration of the 13C label into the plant–soil system.

2.3. Soil and plant sampling

Sampling of soil and plants was carried out prior to (control) and24 h after (labeled) the pulse-labeling. Control samples were takenat approximately 1 m distance from the selected pulse-labelingarea. Soil samples were taken from 0 to 5 cm depth and were kepton ice in the field, transported to the laboratory and stored ina �24 �C freezer.

2.4. Soil and plant processing and measurements

The soil samples (both labeled and control) were thawed andsieved moist through a 2 mm sieve to remove all visible roots,macro fauna and fresh litter since plants contain large concentra-tions of the universal PLFA 16:0, as well as PLFAs 18:2u6,9 and18:3u3,6,9 (Harwood and Russel, 1984; Stahl and Klug, 1996; Zelles,1997) which are generally used as biomarkers for fungi. Duringsieving, roots were removed from the soil by forceps and washedwith water to separate the root-adhering soil from the roots. Boththe bulk (BU) and root-adhering (RA) soil were freeze-dried andstored in the freezer until further processing. Root and shootbiomass was dried at 45 �C for 48 h and stored dry until furtheranalysis. Sub-samples of the freeze-dried soil and oven dried plantmaterials were ground and analyzed in duplicate on an ElementalAnalyzer (ANCA-SL, PDZ-Europa, UK) coupled to an Isotope RatioMass Spectrometer (model 20-20, SerCon, UK) for total C, total Nand 13C content. Carbon isotope ratios (d13C) were expressed permil (&), and are the relative measurement against the 13C/12C ratioin an international standard (Vienna Pee Dee Belemnite (VPDB)):

d13C ¼Rsample � Rstandard

Rstandard� 1000ð&Þ

with R¼ 13C/12C. The working standard was flour with a carboncontent of 39.95% and d13C value of �27.01& vs. VPDB (both valuescertified by Iso-Analytical, UK).

2.5. PLFA extraction, quantification and 13C analysis

The extraction, quantification and compound-specific 13C anal-ysis of phospholipid fatty acids (PLFAs) were performed followingthe method described by Denef et al. (2007). Total lipids wereextracted from 6 g of freeze-dried soil using phosphate buffer/chloroform/methanol at a 0.9:1:2 ratio and partitioned into neutral,glyco- and phospho-lipids by solid phase extraction. Phospholipidswere methylated by mild alkaline methanolysis (using methanolicKOH) to form fatty acid methyl esters (FAMEs), which wereanalyzed by capillary gas chromatography–combustion-isotoperatio mass spectrometry (GC–c-IRMS) (GC-C/DeltaPLUS XP ThermoScientific) via a GC/C III interface as described by Denef et al. (2007).On average, 25 PLFA peaks were detected, but only 16 were selectedfor 13C analysis because of their use as biomarker fatty acids fordifferent microbial communities (Zelles, 1997). These PLFAscomprised approximately 90% of the total PLFA concentration. ThePLFAs i-C15:0, a-C15:0, i-C16:0, i-C17:0, a-C17:0 were designated asgram-positive bacterial whereas cy-C17:0, cy-C19:0, C16:1u7 andC18:1u7 were designated as gram-negative bacterial (Frostegårdand Bååth, 1996; Zelles, 1997). C18:1u9, C18:2u6,9 and C18:3u3,6,9were used as indicators of saprophytic fungi (Frostegård and Bååth,1996; Stahl and Klug, 1996; Zelles, 1997), while C16:1u5 was usedto indicate arbuscular mycorrhizal fungi (Olsson, 1999). The PLFAs10Me-C16:0 and 10Me-C18:0 were used to indicate soil actino-mycetes (Kroppenstedt et al., 1984; Brennan, 1988). The universal

PLFA C16:0, occurring in the membranes of all organisms, wasgenerally the most abundant PLFA. Standard fatty acid nomencla-ture was used to describe PLFAs. The number before the colon refersto the total number of C atoms; the numbers following the colonrefer to the number of double bonds and their location (after the‘u’) in the fatty acid molecule. The prefixes ‘‘Me,’’ ‘‘cy,’’ ‘‘i,’’ and ‘‘a’’refer to the methyl group, cyclopropane groups, and iso- andanteiso-branched fatty acids, respectively. Most biomarker PLFAswere completely separated using a CP-SIL88 GC-column(100 m� 0.25 mm i.d.� 0.2 mm film thickness; Varian Inc.), exceptfor the PLFAs C16:1u7 and a-C17:0 which occasionally merged intoone peak (designated C16:1u7þ). In the chromatograms whereC16:1u7 and a-C17:0 were fully separated, a much greater peakarea was observed for C16:1u7, suggesting a dominant gram-negative bacterial origin.

For each sample, the abundance of individual PLFAs was calcu-lated in absolute amounts of C (nmol PLFA-C g�1 soil) and thenconverted to mole percentage PLFA-C. The d13C values of the indi-vidual FAMEs obtained from the GC–c-IRMS were corrected for theaddition of the methyl group during transesterification by simplemass balance:

d13CPLFA ¼½ðNPLFA þ 1Þd13CFAME � d13CMeOH�

NPLFA

where NPLFA refers to the number of C atoms of the PLFA compo-nent, d13CFAME is the d13C value of the FAME after transesterification,and d13CMeOH is the d13C value of the methanol used for trans-esterification (�36.7� 0.4& vs. VPDB by EA-IRMS using Chromo-sorb-W; n¼ 20).

2.6. Statistical analyses

All statistical analyses were computed using the program R 2.4.1(The R Foundation for Statistical Computing, 2006). All data werea priori tested for normality by Kolmogorov–Smirnov test in the‘nortest’-package, analyses of variance were executed in the ‘stats’-package and principal component analysis in the ‘ade4’-package.

The effects of soil habitat and mowing frequency were testedsingularly, by one-way analysis of variance, on the mole PLFA-Cpercentage and 13C enrichment, for each biomarker PLFA. Forexample, the effect of soil habitat was tested within each singlecombination of mowing frequency and nitrogen fertilization. In thisway the influence of other treatments, even when found non-significant, was excluded. The effect of N-fertilization was tested onpooled data of mowing treatments for biomarker PLFAs that did nothave a significant mean difference between the 3M and 5M treat-ments. This pre-planned approach was based on analysis of vari-ance in soil properties and abundance of biomarker PLFA betweenthe mowing frequency treatments. The threshold confidence levelfor significance testing was set at 5%, meaning the null-hypothesiswas rejected when the probability was smaller than 0.05 and theeffect of the treatment was considered significant. The significanceof difference between the three N-fertilization levels was a poste-riori tested by computation of least square differences (LSD). MolePLFA-C percentages were standardized for all biomarker PLFAsprior to statistical analyses since normality assumptions were notmet for a few biomarker PLFAs.

Principal component analysis (PCA) on the mole percentages ofbiomarker PLFA was used to identify the most discriminatoryeffects on the abundance of microbial groups. Standardization ofthe mole percentages was necessary to meet normality assump-tions. The percentage of the variance explained by a treatmentwithin the complete dataset was determined as the fraction of thevariance within a treatment to the variance in the complete dataset.The significance of differences between PLFA profiles was tested by

Fig. 1. Total PLFA-C (nmol PLFA-C g�1 soil) in bulk (BU) and root-adhering (RA) soilsamples taken after pulse-labeling. Lower case letters and asterisks indicate thesignificance of the difference between soil habitats (*: P< 0.05, **: P< 0.01 and ***:P< 0.001 by LSD post-hoc test, n¼ 4). No significant differences were found between Nfertilizer treatments or between mowing treatments.

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153 147

a multi-response permutation procedure (MRPP); this algorithmcalculates all pair wise distances in the entire dataset (Mielke andBerry, 2001). The Euclidean measure of distance was used in thisanalysis to be compatible with PCA. MRPP provides a change-cor-rected within-group agreement: A¼ 1� delta/E(delta), where deltais the overall weighted mean of group mean distances and E(delta)is the expected delta assessed as the average of the permutatedsampling units. The significance test is the fraction of the permuteddeltas that are less than the observed delta.

3. Results

3.1. Soil and plant properties

Soil C and N content, soil C/N ratio and soil pH did not signifi-cantly differ among N fertilizer and mowing treatments (Table 2).However, the C/N ratio of plant shoots and roots significantlydecreased with increased N addition. Increased mowing frequencyalso decreased the C/N ratio of plant shoots, but only in the 225Ntreatment.

3.2. Phospholipid fatty acid analyses

The total concentration of PLFA-C did not significantly differamong N fertilizer or mowing treatments. In the N-fertilizedtreatments, total PLFA-C concentrations were always greater in theRA than in the BU soil habitat, but this was only significant in the450N/5M treatment (Fig. 1).

The mole percentages of the biomarker PLFAs investigated inthis study are shown in Table 3 for the BU soil (Table 3a) and RA soil(Table 3b). The highest mole percentage was found for the universalbiomarker PLFA C16:0 in both BU and RA soil samples, while thefungal biomarker C18:3u3,6,9 was relatively the least abundant.Fig. 2 shows the plots of the PCA based on the PLFA profile of themicrobial community (i.e. mol% PLFA-C). Within the completedataset, soil habitat was the most discriminatory factor on the PLFAprofiles, accounting for 26% of the variance in PLFA profiles basedon between-group analysis. The first principal component (PC1) inthe PCA analysis, summarizing 35% of the variance in the completedataset, distinguished the RA from the BU habitat. There wasa significant difference in PLFA profiles between both soil habitats,as indicated by MRPP analysis. The effect of N fertilization andmowing frequency were therefore investigated per soil habitat.

Mowing treatment only significantly affected the relativeabundance of two biomarker PLFAs in the 450N treatment (Table3): in the RA soil, the proportion of C16:1u7þ decreased withincreased mowing frequency, while in the BU soil, the proportion of

Table 2Means (n¼ 4) of plant- and soil-related properties in the different grassland treat-ment plots. Soil analyses were done on 2 mm sieved control soil, after manual rootremoval.

Properties 0N/3M 225N/3M 225N/5M 450N/3M 450N/5M

Soil %C 1.28 1.77 1.51 1.24 1.14Soil %N 0.12 0.16 0.14 0.12 0.11Soil pHa 5.2 4.6 4.5 3.9 4.1Soil C/N 11.1 11.3 11.0 10.4 10.8Shoot C/N 49.1 a 36.3 bA 24.5 dB 20.7 c 15.0 eRoot C/N 38.3 a 30.9 b 27.6 d 21.1 c 21.5 e

For a given parameter, N fertilizer treatment means within a mowing treatmentfollowed by different lower case letters differ significantly (P< 0.05 by LSD test,n¼ 4). For a given parameter, mowing treatment means within an N fertilizertreatment followed by different upper case letters differ significantly (P< 0.05 byLSD test, n¼ 4).

a Soil pH was determined on unlabeled soil samples in 0.01 M CaCl2 (1:5 soil:-ClaCl2 suspension).

10Me-C16:0 increased with increased mowing frequency. Nitrogenaddition increased the relative abundance of several gram-positivebacterial PLFAs (a-C15:0; i-C16:0, i-C17:0), gram-negative bacterialPLFA C17:0cy and non-specific PLFA C18:0 in the BU soil (Table 3).However, N addition reduced the mole percentage of C18:1u7(gram-negative biomarker) in BU and RA and C16:1u5 (AMFbiomarker) in RA (Table 3).

3.3. 13C incorporation into soil, shoots and roots

The d13C values of plant shoots, plant roots, bulk (BU) soil androot-adhering (RA) soil before (control) and 24 h after (labeled)pulse-labeling are presented in Table 4. For control samples, BU andRA soil samples were generally less depleted in 13C compared toplant shoots and roots. After pulse-labeling, the highest enrichmentin 13C was found in the shoots followed by the roots, while the BUand RA soil had the lowest 13C enrichment. High variability in the13C incorporation into soil, shoot and root samples was foundamong replicates. Although 13CO2 supply differed among treat-ments during pulse-labeling due to different CO2 uptake rates, 13Cenrichment of aboveground plant biomass was quite comparableacross treatments (Table 4). 13C enrichment of plant roots on theother hand differed more among treatments, with the greatestenrichment in the 0N/3M treatment and the lowest in the 450N/5Mtreatment.

3.4. 13C incorporation into microbial PLFAs

The d13C values of PLFAs in the control soils varied amongindividual PLFAs and ranged between �22.3& and �34.4& (datanot shown). Occasionally, very depleted d13C values as low as�50&

were obtained for C18:3u3,6,9, a commonly used fungal biomarkerPLFA but also present in plants. Pulse-labeling with 13CO2 resultedin an increase in the d13C value of most identified biomarker PLFAs(Table 5). Although associated with high variability, post-labelingd13C values differed greatly among individual PLFAs, suggestingdifferent uptake rates of new rhizodeposit-C by different microbialcommunities. In the non-fertilized treatment, the highest d13C labelwas found in fungal biomarkers C18:1u9, C18:2u6,9, C18:3u3,6,9

Table 3Mean relative abundance (mol PLFA-C %) and standard deviation (in brackets) (n¼ 4) of individual biomarker PLFAs in bulk soil (a) and root-adhering soil (b).

Community PLFA 0N/3M 225N/3M 225N/5M 450N/3M 450N/5M

(a) Bulk soilActinomycetes 10Me-C16:0 3.00 (0.37) a 4.12 (0.24) b 4.55 (0.88) b 3.81 (0.71) Ab 5.62 (0.89) B

10Me-C18:0 2.77 (0.42) a 2.70 (0.23) ab 2.29 (0.44) ab 2.03 (0.31) b 1.79 (0.29) b

Fungi C18:1u9 7.08 (2.74) 5.92 (0.14) 6.31 (0.46) 5.92 (0.59) 5.35 (0.46)C18:2u6,9 8.80 (0.70) 10.60 (2.19) 11.35 (3.26) 11.28 (1.77) 9.44 (1.31)C18:3u3,6,9 0.93 (0.34) 0.95 (0.31) 1.10 (0.61) 1.12 (0.37) 0.70 (0.01)

AM Fungi C16:1u5 5.02 (0.57) a 2.76 (0.57) b 3.01 (1.32) b 2.92 (0.70) b 4.94 (2.91) b

Gram-negative bacteria C16:1u7þ 6.84 (1.47) 7.16 (1.45) 6.82 (1.91) 6.60 (1.84) 6.94 (0.63)C17:0cy 2.58 (0.26) a 3.15 (0.33) b 3.16 (0.18) b 3.89 (0.36) b 3.62 (0.47) bC18:1u7 10.48 (0.47) a 6.98 (0.91) b 7.07 (1.07) b 5.05 (0.76) c 4.62 (0.62) c

Gram-positive bacteria i-C15:0 9.35 (0.66) 9.58 (0.64) 9.32 (1.21) 8.66 (0.59) 9.29 (0.71)a-C15:0 5.62 (0.40) a 6.70 (0.46) b 6.27 (0.71) b 6.44 (0.32) ab 6.24 (0.44) abi-C16:0 3.25 (0.08) a 6.30 (0.42) b 5.88 (0.72) b 5.85 (0.34) b 6.01 (0.59) bi-C17:0 2.85 (0.27) a 3.26 (0.31) ab 3.25 (0.31) ab 4.20 (0.78) b 4.72 (0.71) b

Non-specific C15:0 0.61 (0.10) 0.69 (0.09) 0.71 (0.11) 0.56 (0.10) 0.64 (0.12)C16:0 18.04 (1.37) 16.73 (1.00) 18.64 (1.66) 18.66 (1.58) 18.39 (1.31)C18:0 3.64 (0.50) a 3.83 (0.62) a 2.88 (0.69) a 6.00 (1.00) b 5.90 (1.62) b

(b) Root-adhering soilActinomycetes 10Me-C16:0 2.58 (0.96) 2.61 (0.69) 3.11 (0.70) 2.89 (1.08) 3.88 (0.52)

10Me-C18:0 3.10 (0.35) 3.04 (0.33) 3.38 (1.11) 2.76 (0.26) 3.65 (0.51)

Fungi C18:1u9c 6.94 (1.25) 7.60 (1.43) 7.29 (0.46) 7.18 (1.41) 8.04 (0.69)C18:2u6,9 5.17 (0.23) 6.94 (1.68) 6.49 (2.86) 6.05 (1.37) 5.03 (0.73)C18:3u3,6,9 0.99 (0.34) 0.50 (0.13) 0.88 (0.33) 1.21 (0.76) 0.62 (0.53)

AM Fungi C16:1u5 4.83 (0.46) 2.44 (0.95) 3.84 (1.25) 3.18 (0.60) 3.14 (0.78)

Gram-negative bacteria C16:1u7þ 7.60 (0.39) 7.69 (2.47) 9.43 (1.14) 9.79 (0.40) A 8.63 (0.40) BC17:0cy 2.21 (0.27) 2.60 (0.78) 2.84 (0.09) 3.32 (0.14) 3.29 (0.44)C18:1u7 10.87 (1.72) a 7.64 (2.56) ab 8.67 (1.46) ab 6.36 (1.26) b 6.77 (1.29) b

Gram-positive bacteria i-C15:0 8.56 (1.26) 8.09 (2.71) 9.37 (0.71) 9.00 (1.22) 9.45 (1.58)a-C15:0 5.17 (0.75) 5.74 (1.87) 6.58 (0.42) 6.85 (0.48) 6.65 (1.01)i-C16:0 3.56 (0.39) 5.22 (1.94) 5.90 (0.23) 5.82 (0.79) 6.49 (0.63)i-C17:0 2.05 (1.35) 2.40 (0.53) 2.79 (0.20) 3.74 (0.49) 3.67 (0.33)

Non-specific C15:0 0.96 (0.18) 1.16 (0.20) 1.03 (0.15) 0.97 (0.10) 1.07 (0.14)C16:0 18.00 (3.87) 21.33 (7.94) 18.46 (1.93) 19.23 (1.59) 18.06 (1.05)C18:0 2.97 (0.39) 8.98 (9.21) 3.53 (0.46) 5.49 (1.00) 4.82 (0.53)

For a given PLFA, lower case characters indicate significant mean differences between N fertilizers treatments and upper case characters significant mean differences betweenmowing treatments within an N fertilizer treatment (P< 0.05 by LSD test after standardization, n¼ 4). The effect of N-fertilization was tested on the pooled data of mowingtreatments for biomarker PLFA that showed no significant mowing effect.

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153148

(saprotrophic fungal biomarkers) and C16:1u5 (arbuscular mycor-rhizal fungal biomarker), with d13C values reaching as high as3000& (C18:3u3,6,9) and 1700& (C18:2u6,9). The lowest 13Cenrichment was found in the PLFAs 10Me-C16:0 and 10Me-C18:0(indicative of actinomycetes) and gram-negative bacterial PLFAC17:0cy. In the BU soil, gram-positive bacterial PLFAs a-C15:0, i-C15:0, i-C16:0 and i-C17:0 had also very low 13C enrichment. Thesegram-positive bacterial PLFAs were more enriched in 13C in the RAsoil compared to the BU soil (Table 5, Fig. 3). Gram-negativebacterial biomarkers C16:1u7þ and C18:1u7 were more enrichedin 13C than gram-positive biomarkers.

Except for the greater 13C enrichment of the gram-positivebacterial PLFAs in the RA compared to the BU soil (Fig. 3), d13Cvalues of all other PLFAs did not significantly differ between soilhabitats. d13C values of individual PLFAs did also not differ betweenmowing treatments for a given N fertilizer treatment. However, Nfertilizer treatment had a significant effect on the d13C value ofseveral PLFAs (Table 5, Fig. 4). The non-fertilized treatment hadgreater 13C enrichment of saprotrophic fungal biomarkers C18:1u9(BU and RA), C18:2u6,9 and C18:3u3,6,9 (BU), AMF biomarker PLFAC16:1u5 (BU and RA), gram-negative bacterial PLFA C18:1u7þ (RA)and non-specific PLFAs C16:0 and C18:0 (BU and RA) compared tothe N fertilized treatments (Table 5, Fig. 4). No major N treatmenteffects could be detected on the 13C enrichment of the

gram-positive bacterial and actinomycetes PLFAs, which becameonly minimally labeled with 13C after pulse-labeling.

4. Discussion

4.1. Rhizodeposit-C assimilating communities

The first objective of our study was to identify the microbialcommunities that are utilizing new rhizodeposit-C in this type ofgrassland ecosystem. In the non-fertilized treatment, PLFAs indic-ative of both saprotrophic as well as AMF fungi appeared to be themost 13C enriched biomarker PLFAs in both soil habitats followingpulse-labeling. On the other hand, the biomarker PLFAs indicativeof gram-positive bacteria and actinomycetes were the leastenriched in 13C following pulse-labeling and this in all treatments.Our data therefore suggest that fungal populations are dominantlyinvolved in the immediate processing of newly deposited root-derived C in these grassland soils, while gram-positive bacteria andactinomycetes prefer other C sources for their growth. The high 13Clabel of C18:3u3,6,9, reaching d13C values as high as 3000&, wasremarkable considering the fact that this PLFA was the leastabundant of all investigated PLFAs in this study (Table 3). As formost grassland PLFA studies, the potential contribution of newlyformed root hairs and sloughed root cells to the abundance of

Fig. 2. PCA ordination based on mole percentages PLFA-C of individual PLFAs from 0–5 cm soil samples; the first and second principle components are depicted. Percentageof variance explained by each component is indicated within parenthesis on each axis.The replicates for the different soil habitat, N and mowing treatments are indicated bydifferent symbols. The significance (P-value) of the difference in mol% PLFA-C of allconsidered biomarker PLFAs between soil habitats, as revealed by MRPP, is noted onthe figure. A¼measure for within-group agreement, between 0 and 1.

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153 149

polyunsaturated PLFAs, such as C18:2u6,9 and C18:3u3,6,9 needsto be considered (Harwood and Russel, 1984; Zelles, 1997), evenafter thorough sieving and manual removal of roots prior to PLFAanalysis.

In other grassland studies, these fungal biomarker PLFAs alsoshowed the greatest incorporation of 13C label immediatelyfollowing in situ 13CO2 pulse-labeling (Butler et al., 2003; Treoniset al., 2004; Denef et al., 2007), or following 13C substrate additionin laboratory incubations (McMahon et al., 2005; Williams et al.,2006), with a much slower uptake of 13C by bacteria and actino-mycetes (Denef et al., 2007). Based on shifts in d13C amongbiomarker PLFAs over time following an in situ grassland pulse-labeling, Denef et al. (2007) postulated a possible fungi-mediatedtransfer of rhizosphere-C to bacterial communities from degener-ating or C-leaking fungal hyphae, or a preferential bacterial use ofdead root or fungal necromass materials as C source over the directutilization of fresh root-exudate C.

4.2. Effect of grassland management on microbial communitycomposition

Many studies have demonstrated the influence of mineralfertilizer addition on microbial biomass and/or activity, and bothdeleterious (e.g. Soderberg and Bååth, 2004; Bittman et al., 2005;

Table 4Mean d13C (&) and standard deviation (in brackets) (n¼ 4) of plant shoot and root biomasslabeling (control) and 24 h following pulse-labeling (labeled).

0N/3M 225N/3M

ControlShoots �28.6 (0.5) �27.9 (0.4)Roots �29.5 (0.5) �28.9 (0.4)RA soil �26.9 (0.2) �27.4 (0.4)BU soil �27.3 (0.4) �27.4 (0.6)

LabeledShoots 1134.5 (191.8) 1840.7 (515.5)Roots 170.9 (68.4) 48.9 (28.4)RA soil �10.1 (8.6) �17.8 (1.9)BU soil �9.5 (2.5) �18.0 (6.3)

Wallenstein et al., 2006) and beneficial (Simek et al., 1999) effectshave been described. Other studies could not find any effect offertilizer addition on microbial indices (Hatch et al., 2000; Peacocket al., 2001; Liu et al., 2007). The effects of fertilizer managementmay depend on the nature and quantity of fertilizer added, initialsoil fertility status, and duration of fertilization (Bardgett et al.,1999a). For example, in a lowland grassland soil, Lovell et al. (1995)found no effect of N addition on the size of the microbial biomassafter 1 year, but in the longer term (10 years) a smaller but moreactive biomass developed in response to regular inputs of N. Fiveyears of N fertilizer application increased the total abundance ofPLFA in our grassland soils, although the differences were onlymarginally significant and this only in the 3M mowing treatment(P¼ 0.06). However, total PLFA content in our study was signifi-cantly correlated with total carbon content of the soil (r¼ 0.91,P¼ 0.032 for RA; r¼ 0.96, P¼ 0.009 for BU) as well as with potentialcarbon mineralization rate (r¼ 0.95, P¼ 0.015). The latter wasdetermined by measuring respiration rates during a 10 day incu-bation of BU samples under optimal temperature and moistureconditions (data not shown). Differences in total PLFA abundancebetween the N fertilizer treatments consequently disappearedwhen correcting for total soil C content. These observations indicatea tight coupling of microbial biomass to carbon availability and arein agreement with findings by Hatch et al. (2000) where fertilizer Ninputs modified the microbial biomass only when there was a cor-responding and significant increase in soil C.

Five years of N fertilization also significantly altered thecomposition of the soil microbial community in these grasslandsoils. Many studies have reported N fertilizer addition effects on themicrobial community composition in terms of a decrease in theratio of 18:2u6,9 to bacterial PLFA (Bardgett et al., 1996, 1999a; deVries et al., 2006), which suggests that fungi play a relatively moreimportant role under less fertilized conditions. However, stimu-lated fungal activity has also been observed upon mineral N addi-tion (Rousk and Bååth, 2007). In our study, the relative abundanceof saprotrophic fungal PLFAs was not affected by mineral N addi-tion. Both direct and indirect effects of fertilizer addition onmicrobial communities have been proposed by others, e.g. throughthe effect on enzyme activity or buildup of recalcitrant and toxiccompounds (Fog, 1988), changes in plant productivity and speciescomposition (Bardgett et al., 1999b; Donnison et al., 2000), soil pH(Bååth and Anderson, 2003; de Vries et al., 2006), trophic structureof the soil fauna (Bardgett et al., 1998), and amount or quality ofplant-C inputs (Mawdsley and Bardgett, 1997; Grayston et al.,2001). In the study by Rousk and Bååth, fungi favored high C/Nsubstrate over low C/N substrate. Fungi are capable of colonizingnutrient-poor and recalcitrant substrates due to their greater andmore variable C/N ratio, their wide-ranging enzymatic capabilitiesand their ability to allocate essential nutrients through theirhyphae over considerable distances (Frey et al., 2003). In our study,5 years of N addition had caused a decrease in the C/N ratio of the

samples, and root-adhering (RA) and bulk (BU) soil of each treatment prior to pulse-

225N/5M 450N/3M 450N/5M

�28.9 (0.8) �28.4 (1.2) �29.6 (0.7)�29.3 (0.2) �29.2 (0.7) �29.8 (0.3)�27.8 (0.6) �26.8 (0.3) �27.5 (0.6)�27.6 (0.5) �27.7 (0.3) �27.8 (0.4)

1239.0 (484.8) 1790.1 (917.8) 1536.1 (389.7)56.9 (57.0) 90.5 (168.7) 0.15 (22.3)�12.1 (6.4) �8.3 (26.4) �22.3 (1.8)�17.0 (7.2) �4.0 (29.9) �22.7 (0.6)

Table 5Mean post-labeling d13C values (&) and standard deviation (in brackets) (n¼ 4) of individual PLFAs in bulk soil (a) and root-adhering soil (b).

Community PLFA 0N/3M 225N/3M 225N/5M 450N/3M 450N/5M

(a) Bulk soilActinomycetes 10Me-C16:0 �25.5 (0.0) �2.8 (27.8) �6.9 (21.1) �27.5 (4.7) �29.1 (6.6)

10Me-C18:0 �27.7 (0.5) �28.7 (3.5) �30.2 (5.4) �23.0 (4.1) �27.1 (3.6)

Fungi C18:1u9 257.9 (91.9) a 103.3 (119.4) b 60.8 (53.1) b 73.1 (75.8) b 33.5 (30.7) bC18:2u6,9 758.3 (317.9) 536.7 (761.0) 400.3 (440.7) 242.6 (150.2) 108.6 (106.3)C18:3u3,6,9 1538.8 (976.9) 1199.2 (1540.3) 548.9 (536.0) 379.4 (174.1) 849.7 (672.5)

AM Fungi C16:1u5 279.6 (90.7) a L20.1 (6.1) b 71.1 (108.6) b L6.2 (15.7) b L21.6 (0.2) b

Gram-negative bacteria C16:1u7þ 68.3 (23.8) 13.0 (5.0) 29.3 (20.4) 45.2 (81.8) 0.7 (17.1)C17:0cy �28.7 (3.5) �27.8 (2.0) �25.7 (4.3) �23.7 (4.0) �26.5 (2.7)C18:1u7 125.0 (19.5) 76.2 (42.6) 94.2 (47.7) 88.5 (92.5) 18.4 (25.8)

Gram-positive bacteria a-C15:0 �9.4 (10.0) �9.7 (8.5) �6.2 (18.7) �1.9 (21.9) �19.2 (1.7)i-C15:0 �6.0 (9.2) �9.8 (9.0) �9.3 (14.9) �5.2 (19.6) �19.6 (2.3)i-C16:0 1.5 (13.3) �7.9 (10.6) �3.2 (21.9) 4.5 (28.5) �15.7 (1.5)i-C17:0 �14.0 (8.5) �12.9 (7.2) �9.2 (17.3) �8.3 (15.4) �18.9 (0.8)

Non-specific C15:0 24.8 (29.2) �1.3 (17.1) �0.4 (20.0) 7.4 (40.6) �26.1 (16.7)C16:0 460.1 (118.9) a 244.5 (317.2) ab 226.6 (216.2) ab 122.9 (93.1) b 52.0 (61.3) bC18:0 255.0 (171.7) a 13.8 (16.7) b 21.7 (36.9) b 9.9 (37.6) b 10.4 (32.2) b

(b) Root-adhering soilActinomycetes 10Me-C16:0 �28.3 (75.3) �15.1 (3.4) �5.2 (10.1) �8.1 (25.0) �18.8 (8.5)

10Me-C18:0 �29.8 (6.0) �27.4 (4.5) �24.5 (4.2) �20.9 (4.9) �24.1 (2.7)

Fungi C18:1u9c 260.2 (134.4) a 29.5 (12.4) b 54.2 (46.5) b 75.4 (142.0) b 28.6 (36.7) bC18:2u6,9 521.7 (255.6) a 62.2 (32.1) b 242.9 (135.2) b 194.1 (270.5) b 101.6 (70.2) bC18:3u3,6,9 644.4 (413.8) a 160.9 (127.4) b 242.4 (140.9) b 150.4 (176.8) b 265.3 (275.3) b

AM Fungi C16:1u5 294.3 (127.3) a L13.3 (1.3) b 89.4 (114.2) b L10.5 (15.1) b L18.8 (3.6) b

Gram-negative bacteria C16:1u7þ 111.2 (54.6) a 20.8 (9.6) ab 54.1 (40.0) ab 28.6 (62.4) b 8.7 (17.0) bC17:0cy �38.3 (18.3) �24.0 (5.2) �25.2 (0.9) �28.7 (2.3) �24.7 (8.5)C18:1u7 191.2 (48.7) 73.3 (27.0) 120.8 (68.7) 100.1 (139.1) 38.1 (36.0)

Gram-positive bacteria a-C15:0 25.3 (44.6) �2.0 (16.5) 14.5 (35.4) 19.9 (61.2) �11.8 (5.3)i-C15:0 21.0 (31.3) �4.5 (12.6) 8.8 (27.7) 16.1 (54.8) �11.5 (4.9)i-C16:0 31.7 (37.6) 0.0 (16.5) 19.3 (34.8) 31.5 (75.2) �7.0 (5.5)i-C17:0 57.1 (95.2) �5.0 (11.7) 16.3 (41.5) 8.0 (44.0) �10.8 (2.9)

Non-specific C15:0 20.5 (5.7) �5.1 (17.6) 10.8 (35.4) 19.8 (62.1) �13.7 (7.4)C16:0 386.9 (118.1) a 42.3 (30.0) b 143.3 (82.4) b 88.6 (147.3) b 37.4 (34.3) bC18:0 355.6 (334.8) a 8.3 (33.9) b 47.5 (54.5) b 23.6 (70.3) b 20.8 (37.9) b

For a given PLFA, N fertilizer treatment means followed by different lower case letters differ significantly (P< 0.05 by LSD test, n¼ 4). The effect of N-fertilization was tested onthe pooled data of mowing treatments, since mowing had no significant effect.

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153150

grass roots without altering the relative abundance of saprotrophicfungal PLFAs C18:1u9, C18:2u6,9 or C18:3u3,6,9. This suggests thatroot quality was not a controlling factor for the relative abundanceof saprotrophic fungi in the microbial community. However,a decrease in the relative abundance of the arbuscular mycorrhizalbiomarker PLFA C16:1u5 was found as a result of N fertilization inthe root-adhering soil. Bradley et al. (2006) showed a decrease inthe proportion of arbuscular mycorrhizae upon short-term Nfertilizer addition while saprotrophic fungal communities wereonly affected in their abundance by long-term N fertilization (18years). However, in forest soils along a natural fertility gradient,Hogberg et al. (2003) noticed an increase in the abundance ofmycorrhizal marker C16:1u5 with higher N availability. In theirstudy, increased N availability was accompanied by an increase insoil pH, while our soils and those of Bradley’s study became moreacidic upon N fertilizer addition. As suggested by Bradley et al.(2006), arbuscular mycorrhizae may be more adapted to a highersoil pH. The effect of N availability on microbial communities maytherefore be rather indirect through its effect on soil pH.

We also found contrasting effects of N fertilization on the relativeabundance of bacterial PLFAs. Three gram-positive bacterial biomarkerPLFAs (a-C15:0, i-C16:0 and i-C17:0) showed a significant increase intheir relative abundance with increasing N fertilization in both soilhabitats, while the relative abundance of C18:1u7 (indicative of gram-negative bacteria) was reduced upon N fertilization. This indicates that

changes within the species composition of the bacterial communityalso occurred as a result of N fertilization. Similar responses of gram-positive (increased abundance) and gram-negative (decreased abun-dance) bacterial PLFAs to N fertilization were reported by Peacock et al.(2001) in a corn cultivated soil.

Mowing intensity did not affect total microbial biomass. This isin contrast to several other studies which reported positive effectsof plant defoliation on microbial biomass and/or activity(Mawdsley and Bardgett, 1997; Macdonald et al., 2006). Stimula-tion of microbial biomass upon plant defoliation is likely to bea result of increased exudation of high energy, simple Ccompounds (Paterson and Sim, 2000) or improved root litterquality due to stimulated plant nutrient uptake (Macdonald et al.,2006). However, most studies where plant defoliation stimulatedmicrobial biomass and/or activity investigated changes betweendefoliated and non-defoliated grasslands. This study did notinclude a control site without a mowing treatment; only anincrease in mowing frequency from 3 times to 5 times per yearwas investigated. It is likely that this increase in mowingfrequency did not sufficiently alter root exudation or root litterquality to result in a noticeable difference in the total microbialbiomass. Indeed, we did not observe a difference in the C/N ratioof the roots between the two mowing treatments (Table 2).However, the C/N ratio of the grass shoots in the 225N treatmentdecreased with increased mowing frequency (Table 2), indicating

Fig. 3. Significant soil habitat effects on post-labeling d13C values of gram-positivebacterial PLFAs (i-C15:0, a-C15:0, i-C16:0, i-C17:0) within the 450N/5M treatment.Letters and asterisks indicate the significance of the difference between fertilizationlevels (*: P< 0.05, **: P< 0.01 and ***: P< 0.001, by LSD post-hoc test, n¼ 4).

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153 151

a higher N allocation to the shoots and thus a higher quality ofshoot litter with increased mowing.

Increased mowing had also no significant effect on the molepercentage of most biomarker PLFAs. This is in contrast to otherstudies, reporting microbial community compositional shifts withplant defoliation. Bardgett et al. (1996) reported lower fungal overbacterial biomass ratios in heavily grazed grasslands compared toless grazed grasslands. They concluded that heavy grazing and thusmore severe plant defoliation favors ‘fast cycles’ dominated by

Fig. 4. Significant N treatment effects on post-labeling d13C values of fungal PLFAs C18:1u9, C(C16:0, C18:0), and gram-negative bacterial PLFA C16:1u7þ in bulk (BU) and root-adheritreatments, since mowing had no significant effect. Letters and asterisks indicate the signP< 0.001, by LSD post-hoc test, n¼ 4). Hollow dots indicate outlier replicates within one tr

labile substrates and bacteria, while light grazing supports ‘‘slowcycles’’ dominated by more resistant substrates and fungi. But, thepractice of grazing includes also other factors besides plant defo-liation that could affect microbial communities, for example byaltering N inputs from faecal and urine deposition, as well aschanging soil structure and aerobicity due to animal compaction.

4.3. Effect of grassland management on rhizodeposit-C assimilatingcommunities

Nitrogen fertilization did not only reduce the proportion of AMFPLFA, but also the 13C incorporation in AMF PLFA following pulse-labeling (Table 5, Fig. 4). Lower 13C enrichment with N fertilizer addi-tion was also observed in the roots (Table 4). This suggests that 5 yearsof N fertilization has reduced AMF fungal activity as a result ofa decreased transport of photosynthate-C to roots and reduced Callocation by plants to their arbuscular mycorrhizal symbionts. Aninhibition of arbuscular mycorrhizal fungi by N fertilization has beenreported in other studies based on observations of decreased abun-dance of the biomarker C16:1u5 (e.g. Bradley et al., 2006; Siguenzaet al., 2006) or decreased root colonization by arbuscular mycorrhizalfungi (e.g. Johnson et al., 2003). Results from an earlier pulse-labelingexperiment in nitrogen-limited temperate grasslands (Denef et al.,2007) indirectly suggested increased plant-C allocation to mycorrhizalsymbionts under N limited conditions. In that study, the relative 13Cenrichment of the arbuscular mycorrhizal PLFA increased underelevated CO2. This apparent stimulated mycorrhizal activity and mostlikely resulting in improved nutrient acquisition by mycorrhizaeprovided a valid explanation for the observed increased plantproductivity under elevated CO2, which is rather atypical undernutrient limited conditions (Kammann et al., 2005).

Similar to AMF fungi, the 13C enrichment of the saprotrophicfungal PLFAs also decreased by N fertilizer addition (Table 5, Fig. 4),suggesting suppressed saprotrophic fungal activity in fertilizedcompared to non-fertilized grasslands. So, whereas the impact of Nfertilization on saprotrophic fungi could not be detected in terms oftheir relative abundance, a clear negative impact was revealed in

18:2u6,9, C18:3u3,6,9, arbuscular mycorrhizal fungal PLFA C16:1u5, non-specific PLFAsng (RA) soil. The effect of N-fertilization was tested on the pooled data of mowing

ificance of the difference between fertilization levels (*: P< 0.05, **: P< 0.01 and ***:eatment.

K. Denef et al. / Soil Biology & Biochemistry 41 (2009) 144–153152

terms of their activity. In the study by Bradley et al. (2006), sap-rotrophic fungal communities were only suppressed (in terms oftheir abundance) after long-term (i.e. 18 years) addition of Nfertilizer, while arbuscular mycorrhizal fungi were already affectedafter 1 year of N fertilizer addition. After 18 years of N fertilizeraddition, Bradley et al. (2006) also recorded a difference in plantspecies diversity among N fertilizer treatments. The lack of an‘abundance’ response of saprotrophic fungi to N fertilization at ourexperimental sites might be attributed to limited changes in theplant community after only a period of 5 years of N fertilizeraddition. Several studies have suggested that the effects of fertilizeraddition on soil microbial communities are largely indirect,through a change in the composition of the plant community(Bardgett et al., 1999a, 1999b), due to plant-specific differences inthe release of root exudates and alterations in nutrient competitionbetween plants and rhizosphere microorganisms.

4.4. Variation in microbial community with soil habitat

In the N-amended treatments, root-adhering soil was charac-terized by greater PLFA concentrations than the bulk soil, thoughstatistically significant differences were only found in the 450Ntreatment. PCA analysis revealed significant differences in themicrobial community structure between bulk and root-adheringsoil. The proximity to roots and greater accessibility to exudate-C inthe root-adhering soil may have promoted growth and rhizode-posit-C uptake by distinct microbial communities in this soilhabitat compared to the bulk soil. In the highest N fertilizer andmowing intensity treatment, we observed a higher assimilation of13C by gram-positive bacteria in the vicinity of the root system(Fig. 3). A preference for root-adhering soil or bulk soil could not bedetected for the other rhizodeposit-C assimilating microbialcommunities. Different results were reported by Lu et al. (2007)following a 13CO2 pulse-labeling experiment in rice microcosms. Intheir study, gram-negative and eukaryotic microorganismsappeared to be most actively assimilating root-derived C in therhizosphere of rice plants, whereas gram-positive bacteria weremore active in the bulk soil. The preference of a microbial group fora specific location in the soil matrix to grow and utilize rhizode-posit-C maybe related to the accessibility to root exudates but also,and perhaps more importantly, to the unique physicochemicalconditions (e.g. oxygen level, pore size and particle size distribu-tion, etc.) of the preferred soil habitat (Hattori, 1988; Bolton et al.,1993; Schutter and Dick, 2002).

5. Conclusions

The incorporation of 13C label in biomarker PLFAs in this in situ13CO2 pulse-labeling experiment showed a rapid uptake of newlydeposited plant-derived C by decomposer and AMF fungi anda clear effect of N fertilizer management on the functioning of bothfungal communities. In particular, N fertilization caused a decreasein the assimilation of plant C by both AMF and saprotrophic fungi,while only a decrease in the abundance of AMF fungi was detectedbased on mol% PLFA analyses. While gram-positive bacterialabundance (mol%) was increased with N fertilizer addition, nodifferences could be detected in 13C enrichment among N fertilizertreatments for these PLFAs. The latter is probably a result of the low13C labeling of these PLFAs following pulse-labeling, suggesting thatgram-positive bacteria are less involved in the immediate pro-cessing of new rhizodeposit-C. We therefore conclude that in situ13C pulse-labeling of PLFAs is an effective tool to detect functionalchanges of those microbial communities that are dominantlyinvolved in the immediate processing of new rhizosphere-C. Ourdata also suggested that microbial community composition differswith proximity to plant roots, and that 5 times vs. 3 times mowing

per year in fertilized grasslands does not affect microbial commu-nity structure or functioning.

Acknowledgments

This work was supported by a research grant and postdoctoralfellowship from the Research Foundation of Flanders (FWO -Vlaanderen) to K. Denef. The authors wish to thank Jorin Botte, EricGillis, Katja Van Nieuland and Jan Vermeulen for assistance withpulse-labeling, sample collection and laboratory analyses. We alsothank Ben Leroy (UGent), Angela Kong and Johan Six (UC Davis) forproviding training on PLFA extraction.

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