Chapter 13

The Use of Stable Isotope Labelingand Compound-Specific Analysisof Microbial Phospholipid Fatty Acidsto Quantify the Influences ofRhizodeposition on MicrobialCommunity Structure and Function

Eric PatersonThe James Hutton Institute, Ecological Sciences, UK

13.1 INTRODUCTION

Carbon (C) fluxes at the root–soil interface are centrallyimportant to the biological functioning of soils in anumber of contexts. The microbial biomass is the “eyeof the needle” through which element transformationsand fluxes occur (Jenkinson, 1977)—processes thatare maintained by the energy-rich organic inputs fromvegetation (Paterson, 2003). The input of organic com-pounds to the rhizosphere, via processes collectivelytermed rhizodeposition, is the dominant C-flux into soiland counters losses via mineralization of soil organicmatter (SOM). Therefore, quantifying rhizodeposition, itsutilization by the soil biota and determining the balancebetween its mineralization to CO2, and its stabilizationto recalcitrant or physically protected forms is central tounderstanding the source–sink capacity of soils. This isparticularly the case for perturbed systems displaced fromequilibrium, for example, by land use or environmentalchange. This understanding is important not only in thecontext of feedback responses of soils to the environmentbut also to the health and sustainability of soils, properties

Molecular Microbial Ecology of the Rhizosphere, Volume 1, First Edition. Edited by Frans J. de Bruijn. 2013 John Wiley & Sons, Inc. Published 2013 by John Wiley & Sons, Inc.

that are closely related to maintenance of SOM stocks(Gregory et al., 2009). The rhizodeposition flux into soilconsists of a complex mixture of organic compounds:soluble root metabolites (e.g., sugars, amino acids, andorganic acids; see Chapter 22) that are passively releasedfrom roots and are readily utilizable substrates for soilmicrobial communities (Jones and Darrah, 1994); secretedcompounds with specific functions in soil (e.g., enzymes,phytosiderophores, and signal compounds); and senescedtissues released through root turnover that provide morecomplex substrates for microbial communities (Patersonet al., 2008a). It is known that the quantity and chemicalquality of the rhizodeposition flux varies even betweenclosely related plant types (Germida and Siciliano, 2001)and is strongly affected as a function of plant develop-ment and in response to environmental factors (Paterson,2003; Nguyen, 2003). As soils are generally C-limitedenvironments for microbial activity (Wardle, 1992),rhizodeposition supports increased microbial growth andmicrobially mediated process rates in the rhizosphere, rel-ative to bulk soil (Buyer et al., 2002; Paterson et al., 2009).However, establishing the importance of rhizodeposition

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142 Chapter 13 The Use of Stable Isotope Labeling and Compound-Specific Analysis

in shaping microbial community structure and quantifyingthe consequences for soil processes is not straightforward.

A host of soil physicochemical conditions (pH, waterregimes, organic matter quality, nutrient availability, etc.)are important selective pressures on microbial populations,meaning that although all soils have highly diverse micro-bial communities the structure of these communities issoil specific (Grayston et al., 2004). As a result, the rhi-zosphere microbial community that develops in responseto the rhizodeposition flux from a root extending throughsoil is strongly affected by the starting point for that com-munity (i.e., bulk soil community composition) and theprevailing conditions in specific soils. Therefore, one-to-one relationships between plant and rhizosphere microbialspecies identity are seldom observed in experiments whereplants are grown in different soils (Innes et al., 2004),leading to the suggestion that soil conditions are strongselective influences on microbial community developmentand that plants are of much lesser importance in shap-ing soil communities (Kennedy et al., 2004). However,the observation that plants do not select for rhizospherecommunities with specific species composition does notnecessarily infer that plants do not shape microbial com-munity structure (see Chapter 15), only that the outcomeis also soil dependent. Further, as there is considerablefunctional redundancy within microbial communities, it isentirely possible that although soil type is a strong deter-minant of species composition, plant selection may causesimilar shifts in the functional diversity of microbial com-munities in rhizosphere soil (Waldrop and Firestone, 2004;Paterson et al., 2008a).

Stable isotope (13C) labeling coupled with tracing ofC-flows into components of the soil microbial communityprovides a powerful means to identify microbial groupsactive in mediating specific soil processes (Patersonet al., 2009). Establishing such relationships directly isparticularly important for soil systems, where the inherentcomplexity and interdependence of components and pro-cesses make interpretation of causes and consequencesof community change highly problematic. Nucleic acidstable isotope probing has been applied successfully toidentify microorganisms active in mediating specialistfunctions (Singh et al., 2004; Singleton et al., 2005).However, the application of this approach to study rhi-zosphere plant–soil interactions is difficult for a numberof reasons. Although C-flow from roots to soil can be13C-enriched by growing plants in a 13CO2 atmosphere,it is an expensive undertaking to label plants for a longenough period and with a high enough 13C-enrichmentto make density separation of nucleic acid fractions fromrhizosphere communities potentially viable (Patersonet al., 2009). In addition, the rhizodeposition C-flux is

chemically diverse and contains many compounds utilizedwidely within soil microbial communities, meaning thatroot-derived C does not have the narrow distribution thatsupports discrimination of specific organisms utilizingthe C-source. Finally, pulse-labeling approaches wherehighly enriched 13CO2 is applied for a small proportionof the total plant growth period results in nonuniformlabeling of root-derived C-flow (Thornton et al., 2004).Typically, this is characterized by preferential enrichmentof root exudate compounds and minimal enrichment ofmore complex inputs derived from root turnover andexcretion of secondary metabolites. This is a significantshortcoming of pulse-labeling approaches as it limitstheir interpretation to the fate of only a component ofthe rhizodeposition flux, failing to capture the fate ofmore complex inputs that are only utilized by restrictedfractions of the microbial community.

The limitations described above can be addressed byutilizing a combination of alternative approaches.

1. Phospholipid fatty acid (PLFA) profiles from soilscan be used to characterize microbial communitystructure, and the profiles are sensitive indicators ofcommunity change (Grayston et al., 2004). The majordisadvantage relative to nucleic acid approaches isthat the level of detail on community compositionprovided is very limited. However, unlike othermicrobial biomarkers, PLFA are indicators of livingorganisms (Zelles, 1999) and can identify shiftsin relative abundance of broad microbial groupswithin communities (Paterson et al., 2007). Forisotope tracing studies, PLFA have the significantadvantage that labeled C incorporation into specificPLFA can be quantified extremely sensitively bygas chromatography combustion isotope ratio massspectrometry (GC-C-IRMS).

2. Model exudate mixtures can be applied to soil withsingle components of these mixtures 13C-labeled toallow their utilization by components of the micro-bial community to be determined by isotopic analysisof PLFA compounds (Waldrop and Firestone, 2004;Paterson et al., 2007). While the approach cannot cap-ture the full complexity of the rhizodeposition C-flux,it does allow the specificity of microbial substrate useand concurrent impacts on community structure to bedetermined in the absence of confounding influencesof roots on soil properties (pH, nutrient availability,water status, etc.).

3. Steady-state, continuous 13C-labeling of plants can beachieved using CO2 with a naturally depleted δ13Csignature. Fossil-derived CO2 with a δ13C signature of−35‰ or less is commercially available (13C depleted

13.2 Methods 143

relative to atmospheric CO2 at −8.5‰) and providesan isotopic shift in plant assimilate that can be tracedinto rhizosphere microbial PLFA. This means of 13C-labeling plants is cost-effective, allowing continuouslabeling over extended periods (many months (Pala-cio et al., 2011)). Therefore, plants can be labeledfrom germination to maturity, ensuring completelyuniform labeling of the rhizodeposition C-flux andunbiased quantitative determination of plant-derivedC in soil C-pools.

These approaches were applied to assess the speci-ficity of microbial use of model root exudate compoundsand relationships between substrate use and microbialcommunity development. In addition to application ofmodel exudates, microbial use of root-derived C-flow andrhizosphere community development was investigatedby application of continuous 13C-labeling of plantsand isotopic analysis of microbial PLFA fractions. Wehypothesized that, even in soil with abundant organicmatter, plant-derived C would shape the development ofrhizosphere microbial communities.

13.2 METHODS

Soil was sampled from unimproved, upland (274 m a.s.l.)grassland at Glensaugh, N.E. Scotland. The soil (freelydraining, humus–iron podzol) was sampled to 35 cm(maximum rooting depth at the site) and the humus,eluvial, and illuvial horizons were sieved (<3.35 mm) andcombined. The resulting combined soil had an organic Ccontent of 14.4% and pH (H2O) of 4.6. Full details ofsoil characteristics, preparation, and storage are providedin Paterson et al. (2007).

13.2.1 Model Exudate ExperimentSoil solution samplers (Eijkelkamp, Giesbeek, TheNetherlands) were used as artificial roots to deliver modelexudate solutions to soil. Full details of the microcosmdesign are given in Paterson et al. (2007). In brief,the exudate solution was a mixture of the followingcompounds: glucose, sucrose, fructose, ribose, arabinose,glycine, valine, glutamine, serine, alanine, malic acid,malonic acid, oxalic acid, and fumaric acid. Eachcompound was supplied to the soil at a rate of 0.1 mgC/day via the artificial root. The total C-addition rate wasrepresentative of rhizodeposition from Lolium perenne(Paterson and Sim, 1999). Initially, all microcosms(except controls receiving water only) received identicalexudate solutions for 7 days. Then, for a further 7 days,the microcosms received the same exudate solutions,

except that in one treatment (n = 4) the glucose wasuniformly 13C-labeled, in another the glycine wasuniformly 13C-labeled, and in the final treatment thefumaric acid was uniformly 13C-labeled. Throughout theexperimental period, the soils were maintained at constanttemperature (15 ◦C) and moisture contents were main-tained by misting the soils with sterile deionized waterevery 2 days for the remainder of the experiment (7 days).

Rhizosphere soils associated with the artificialroots were collected (see Paterson et al., 2007), andsubsamples of rhizosphere and bulk soils were frozen at−20 ◦C and freeze-dried before extraction of PLFA(White et al., 1979; Bligh and Dyer, 1959). Aliquotsof PLFA extracts were used for (i) quantification ofPLFA abundance (via standard GC) and (ii) for PLFA13C-enrichment (GC-C-IRMS). Full details of chromatog-raphy conditions, calibration, and quality control aregiven in Paterson et al. (2007).

13.2.2 Continuous 13C PlantLabeling ExperimentThe same soil as used in the model exudate experimentwas packed into Perspex “sheet microcosms.” Half ofthe microcosms (n = 10) were fertilized with ammo-nium nitrate (equivalent to 115 kg N/ha), the remainderreceiving water addition only. Half of the microcosmsin each fertilization treatment (n = 5) were plantedwith four germinated seeds (5–10 mm root emergence)of L. perenne, while the remainder were not planted.Microcosms were transferred to a labeling chamberhoused within a controlled environment room (Conviron,Winnipeg, Canada). The ambient room temperature wasregulated to maintain constant temperature (22 ± 0.5 ◦C)within the labeling chamber during the light (16 h, 500µmol/m/s) and dark (8 h) diurnal cycle. The labelingchamber was continuously flushed (18 l/min) with aircontaining CO2 with a depleted δ13C signature (371 ± 4µmol/mol, −36.1‰ ± 0.2‰). The labeling atmospherewas generated by the removal of CO2 from a compressedair supply and the introduction of CO2 from a cylinderof pure CO2 (BOC, Worsley, UK). Full details of theequipment and experimental design are provided inPaterson et al. (2007). Constant labeling conditions weremaintained during a 28-day growth period, prior todestructively harvesting the systems. At harvest, rhizo-sphere, bulk soil and soil from unplanted microcosmswere collected and analyzed for PLFA concentrations andisotopic enrichments, as described previously.

Full details of associated plant and soil analyses, cal-culations, and statistics applied are detailed in Patersonet al. (2007).

144 Chapter 13 The Use of Stable Isotope Labeling and Compound-Specific Analysis

13.3 RESULTS

13.3.1 Model Exudates ExperimentThe delivery of labeled compounds significantly(P < 0.01) increased the 13C-enrichment of rhizospheresoil (mean across treatments 172.1‰ ± 56.5‰), relativeto bulk soil (−26.8‰ ± 0.1‰). The exudate additionresulted in significant increases in PLFA abundance inrhizosphere soil (125 ± 23 µg PLFA/g soil), relative toaddition of water only (33 ± 8 µg PLFA/g soil). Theexudate addition also significantly increased phosphataseactivity in rhizosphere soil relative to soils receivingwater only (1739 ± 147 and 663 ± 41 µg nitrophenol/h,respectively).

Exudate addition significantly affected the relativeabundance of PLFA biomarkers for microbial groups(Table 13.1), with greater contributions (P < 0.05), inparticular, from gram-negative and fungal markers to thetotal PLFA pool. The increased relative abundance ofthese microbial groups was coincident with high specificincorporation (Inr) of 13C (from each labeled exudatecompound) into their PLFA biomarkers. Where Inr > 1,it indicates that the amount of 13C-incorporation intogroup biomarkers is greater than is accounted for by theirrelative abundance within the whole community.

The distribution of 13C among PLFA compoundswas strongly affected by the chemical source of the13C (Fig. 13.1). For 13C-glucose and 13C-fumaric acid,the 13C was widely distributed among PLFA fractions(25 and 26 PLFA were significantly 13C-enriched,respectively). However, the 13C derived from glycinewas recovered in only nine PLFA compounds. At thelevel of PLFA indicator groups, the distribution of 13Cwas equivalent for glucose and fumaric acid sources.However, for glycine, 13C was recovered to a greaterextent in gram-negative PLFA and to a lesser extent

Figure 13.1 Principal components analysis of 13C distributionamong microbial PLFA fractions from soil around an artificial rootdelivering exudate compounds. Single exudate compounds (glucose,fumarate, or glycine) were replaced with their 13C-labeled analogsduring the treatment period. Source: Modified from Paterson et al.(2007), with permission.

in gram-positive, fungal, and actinomycete fractions(P < 0.05).

13.3.2 Continuous 13C PlantLabeling ExperimentContinuous labeling resulted in a uniform depletionof δ13C in the root and shoot tissues of L. perenne(Table 13.2), relative to unlabeled plants (−26.8‰).However, the depletion of rhizosphere δ13C was notstatistically significant. Principal components analysis(PCA) of microbial community structure (relative abun-dance of PLFA compounds) indicated that rhizospherecommunities were distinct from those in bulk soil and thatN-fertilization resulted in distinct communities in bulk

Table 13.1 Relative abundance of PLFA indicator groups (mol%) in artificial root rhizosphere and bulk soils suppliedwith 15 root exudate compounds and relative incorporation of 13C label from glucose, glycine, or fumaric acid

% Total PLFA % Total Label

Indicator Group Rhizosphere Bulk Glucose Glucose-Inr Glycine Glycine-Inr Fumaric acid Fumaric acid-Inr

Gram negative 28.0 ± 3.5a 15.3 ± 1.1b 41.79 ± 4.6 2.75 ± 0.41b 59.88 ± 6.7 3.91 ± 0.51a 37.83 ± 3.8 2.48 ± 0.31b

Gram positive 10.8 ± 1.1a 7.2 ± 0.4b 8.41 ± 1.7 1.19 ± 0.28c 2.25 ± 0.7 0.32 ± 0.10c 8.42 ± 1.0 1.21 ± 0.29c

Fungi 2.4 ± 0.3a 0.8 ± 0.2b 4.12 ± 0.7 5.19 ± 0.78a 0.90 ± 0.2 1.15 ± 0.08b 3.30 ± 0.56 4.09 ± 0.40a

Actinomycete 3.0 ± 0.1 3.3 ± 0.1 2.12 ± 0.3 0.54 ± 0.08d 0.32 ± 0.6 0.10 ± 0.02c 2.66 ± 0.33 0.82 ± 0.05c

From Paterson et al. (2007), with permission.

The parameter Inr is the net relative incorporation of 13C-label into indicator PLFAs relative to their abundance in bulk soil. Results are means(n = 5) ± standard errors. For % total PLFA, different letters indicate significant differences (P < 0.05) between bulk and rhizosphere abundance ofindicator groups. For Inr, different letters indicate significantly (P < 0.05) different relative incorporation of 13C-label derived from the respectiveexudate compounds into indicator groups.

13.4 Discussion 145

Table 13.2 Lolium perenne growth, partitioning and δ13C of shoot, root, and rhizosphere soil with cultivation inatmospheres containing 13C-depleted CO2 (370 µmol/mol) with and without ammonium nitrate additions to soil

Treatment Plant Weight (mg) Root Weight (mg) Shoot δ13C Root δ13C Rhizosphere δ13C

Unfertilized 53.6 ± 2.1* 33.5 ± 1.9* −52.48 ± 0.42 −52.28 ± 0.33 −27.84 ± 0.05N-fertilized 39.2 ± 2.2* 16.3 ± 1.0* −51.96 ± 0.20 −51.32 ± 0.49 −27.80 ± 0.11

From Paterson et al. (2007), with permission.

Results are means (n = 5) ± standard error, significant (P < 0.05) differences between pairs of means in each column are indicated by “*.”

Figure 13.2 Principal components analysis of PLFA relativeabundance in Glensaugh soil as a function of nitrogen fertilization,planting with Lolium perenne, and location in soil (rhizosphere orbulk). Source: Modified from Paterson et al. (2007), with permission.

soil (Fig. 13.2). However, the rhizosphere communitystructure did not discriminate as a result of N-fertilization.

Although the whole rhizosphere soil was not sig-nificantly 13C-depleted as a consequence of labeling,the depleted δ13C-signature could be readily detected inmicrobial PLFA from rhizosphere soil. Gross patternsof label incorporation were similar across fertilizationtreatments, with the same group of 12 PLFA having thelargest δ13C depletions in each treatment. At a fine level,PCA indicated that the relative fate of plant-derived Cwas quantitatively affected as 13C-fate discriminated onPC2 as a function of fertilization treatment (Fig. 13.3).At the level of PLFA indicative of microbial groups, N-fertilization resulted in a significant (P < 0.05) increase inplant-derived C incorporation into gram-negative bacteriaand reduced incorporation into fungi (Table 13.3).

Figure 13.3 Principal components analysis of the relativedistribution of plant-derived carbon in microbial PLFA fractionsrecovered from the rhizosphere of Lolium perenne, as affected bynitrogen fertilization of the soil. Source: Modified from Patersonet al. (2007), with permission.

13.4 DISCUSSION

13.4.1 Model Exudates ExperimentAddition of model root exudate compounds resulted inincreased microbial PLFA abundance and phosphataseactivity in rhizosphere soil. This general response iscommonly observed for comparisons between plantrhizosphere communities relative to those in bulk soil andconfirms that the exudate addition alleviated microbialgrowth limitation in the experimental systems. Glucose-and fumaric acid-derived 13C was recovered from abroad range of microbial PLFA, consistent with thesecompounds representing substrates readily utilized bymost soil microorganisms (Anderson and Domsch, 1978).The recovery of glycine-derived 13C was restricted tofewer PLFA compounds, suggesting that a relativelyrestricted fraction of the microbial community utilizedthis C-source. Although the physiological basis ofspecificity in microbial use of glycine is unclear, theresult is consistent with a number of previous studies(Lipson et al., 1999; Alef and Kleiner, 1986; Kielland,

146 Chapter 13 The Use of Stable Isotope Labeling and Compound-Specific Analysis

Table 13.3 Relative abundance of PLFA indicator groups in Lolium perenne rhizosphere and bulk soils(± N-fertilization) and fate of plant-derived C in PLFA indicator groups in the rhizosphere

Indicator PLFA Abundances (mol%) Fate of Plant-Derived C (%)

Group NF-Bulk NF-Rhizosphere F-Bulk F-RhizosphereNF-RhizosphereF-Rhizosphere NF-Inr F-Inr

Gramnegative

15.0 ± 1.8* 20.4 ± 2.1* 15.8 ± 1.9* 23.1 ± 2.3* 26.5 ± 4.2* 34.9 ± 5.1* 1.79 ± 0.42a 2.21 ± 0.37a

Grampositive

8.0 ± 0.9* 5.7 ± 0.7* 5.1 ± 0.6 7.6 ± 1.1 4.8 ± 0.8 5.2 ± 0.8 0.58 ± 0.21b 1.07 ± 0.22b

Fungal 0.9 ± 0.1 1.0 ± 0.1 0.3 ± 0.0* 1.1 ± 0.2* 7.7 ± 0.9* 2.3 ± 0.7* 8.57 ± 0.49c 7.77 ± 0.56c

Actinomycete 3.6 ± 0.7 4.0 ± 0.4 3.0 ± 0.3 3.4 ± 0.4 0.4 ± 0.1 0.3 ± 0.1 0.11 ± 0.02d 0.10 ± 0.01d

From Paterson et al. (2007), with permission.

Inr is the net relative incorporation rate of plant-derived C, relative to indicator PLFA abundance in bulk soil. Results are means (n = 5) ± standarderrors. Significant differences (P < 0.05) between means in paired columns are indicated by “*.” Inr is the net relative incorporation of plant-derivedC, relative to indicator PLFA abundance in bulk soil. Significant differences in Inr between PLFA groups, for each treatment, are indicatedby distinct superscript letters.

1995). As glycine has been shown to be a potentiallyimportant direct N-source to plants (Schimel and Chapin,1996), it may be valuable to characterize the importanceof microbial community composition with respect to thecapacity for microbes to compete with plants for glycineacquisition. As a caveat, it should be noted that over the7-day 13C-labeling period, there would have been sometransfer of 13C (e.g., via turnover and predation) fromprimary utilizers to other organisms in soil (Fitter et al.,2005). Therefore, it is possible that slower uptake orreduced turnover of glycine-derived 13C contributed tothe more restricted incorporation of C from this source.These influences could be characterized by time-courseanalysis of 13C-distribution and quantification of 13C-mineralization (13CO2 evolution) rates, as has now beendone for a range of substrates in subsequent experiments(Paterson et al., 2011a, 2011b; Paterson et al., 2008b).

The importance of model exudate compounds, notonly in supporting increased general microbial activity butalso in shaping community structure, is evident from pos-itive relationships between incorporation of 13C into spe-cific PLFA and altered relative abundance in rhizospheresoil (Paterson et al., 2007). Bulk soil community structureis the starting point from which rhizosphere communi-ties develop, influenced by exudate release and other rootprocesses. It was evident that net relative incorporationof exudate C (Inr) was greater for PLFA (both individ-ual compounds and aggregated biomarker groupings) thatincreased in abundance in rhizosphere soil. This resultsuggests strongly that, under the conditions of the experi-ment, the relative ability of microbial populations to com-pete for exudate compounds shaped microbial communitystructure in the rhizosphere. Gram-negative and fungalgroups were those that demonstrated the greatest specificincorporation of 13C (Inr > 1) and also increased theirbiomass fraction within the rhizosphere community to the

greatest extent. It should be noted that this relationshipcould also occur where a factor other than C-availabilitywas limiting, as 13C-uptake would be in proportion to rel-ative activity (imposed by the limiting factor). However,it is well established that C-substrate availability is thedominant microbial limitation in soils (Wardle, 1992), andfor this experiment, the increased microbial biomass sizeand activity following exudate addition demonstrated thatexudate addition alleviated microbial growth limitation.In addition, the artificial root did not impose other poten-tially selective pressures that may be important influencesaround plant roots (e.g., pH and nutrient availability),and delivery of water only via the artificial root did notaffect microbial community structure. Therefore, it is evi-dent that exudate availability and the relative ability ofmicrobial populations to utilize these substrates shapedthe development of community structure.

13.4.2 Continuous 13C PlantLabeling ExperimentThe 13C-labeling with fossil-derived CO2 resulted inδ13C-depletion of plant tissue by approximately 26‰. Asthe labeling was continuous (and steady state) throughoutplant growth, all plant C-pools, and consequently therhizodeposition C-flux, was labeled homogeneously. It isimportant to note that when applying near natural abun-dance labeling (as here), homogeneous labeling cannotbe taken to infer that δ13C values are constant acrossall plant fractions. This is because 13C-fractionation(physical and kinetic processes) results in variationin absolute δ13C-signatures across pools. However,homogeneous labeling ensures that the magnitude of13C-depletion is constant for all pools. Similarly, althoughthe δ13C-signature of soil PLFA fractions was foundto vary between −25‰ and −31‰, in the calculation

References 147

of plant-derived C incorporation this natural abundancevariation is accounted for by measuring the shift in δ13Cduring the experiment rather than only considering theabsolute δ13C-values.

The N-fertilization treatment clearly affected bulk soilmicrobial community structure (Fig. 13.2), notably reduc-ing the abundance of fungi relative to bacteria. This impactof N-availability in soil has been observed as a conse-quence of fertilization of upland grasslands and along tran-sects of natural fertility gradients (Bardgett et al., 1999;Grayston et al., 2001; Brodie et al., 2002; Kennedy et al.,2004). However, although N-fertilization resulted in shiftsin community structure in bulk soil, the correspondingrhizosphere community structures did not discriminate onPCA plots (Fig. 13.2). This demonstrates the selectiveinfluence of rhizosphere conditions, resulting in relativeconvergence of community structures from the distinctcommunities in fertilized and unfertilized bulk soils.

The specific incorporation of plant-derived C intoPLFA was greatest for gram-negative and fungal biomark-ers (Table 13.3), and these groups strongly increased theirrelative abundance within the rhizosphere community.These clear relationships between specific incorporationof plant-C and impacts on relative abundance are consis-tent with results from the model exudates experiment andstrongly support the importance of the rhizodepositionC-flux in shaping microbial community structure.

An advantage of the continuous labeling approachthat has been increasingly exploited since the Patersonet al. (2007) study is that it allows quantitative measure-ment of the balance of plant- and SOM-derived C-useby components of the rhizosphere microbial community.Paterson et al. (2007) found that the proportion ofmicrobial biomass derived from plant-C (quantifiedfrom the δ13C-signature of PLFA compounds) variedwidely among microbial groups (Table 13.4). The highestproportions of biomass derived from plant-C were forgram-negative and fungal groups, with actinomycetebiomass incorporating a much lesser proportion ofplant-derived C. This reflects the relative abilities of thesegroups to access plant- and SOM-derived C-sources.Subsequent studies (Paterson et al., 2008b; Patersonet al., 2011b) have demonstrated that during rhizospherecommunity development, the proportion of plant-derivedC in microbial biomass tends toward a maximum value,in equilibrium with SOM-derived C. These studiesindicate that SOM is a quantitatively important C-sourceto rhizosphere communities and that the balance ofC-source utilization varies among microbial groups.

An important implication of SOM utilization byrhizosphere communities is that the increased microbialactivity around roots not only is associated with utiliza-tion of root-derived C-flow but also stimulates increasedSOM mineralization relative to bulk soil (Paterson

Table 13.4 Effect of nitrogen fertilization on thepercentage of biomass C derived from Lolium perenneassimilate for microbial groups defined by PLFAbiomarker compounds.

Indicator Group Nonfertilized Fertilized

Gram-positive PLFAs 7.56 7.67Gram-negative PLFAs 10.37* 17.08*Actinomycete PLFAs 0.56 0.42Fungal PLFA 48.17* 12.12*

From Paterson et al. (2007), with permission.

Results are means (n = 5) ± standard error, significant differences (P< 0.05) in group utilization of plant-C with N-fertilization areindicated by “*” in each row.

et al., 2008b). This plant-induced “priming” of SOMmineralization and its controls are not well understood(Kuzyakov, 2010), but it has important implications forcycling of nutrients from recalcitrant organic moleculesto forms available to plants (Paterson, 2003). Priming isalso increasingly recognized as a quantitatively significantprocess affecting the dynamics of SOM turnover, but onethat is not accounted for in current soil C models basedon first order kinetics (Ostle et al., 2009; Kuzyakov,2010). An important recent advance in relation to thestudy of priming effects is that the continuous labelingapproach can be coupled with isotopic partitioning of soilrespiration, which allows direct measurement of SOMmineralization in planted soils (Paterson et al., 2008b;Paterson et al., 2011b). Therefore, there are now excellentopportunities to quantify the magnitude of priming effectsand also to probe the microbial mechanisms drivingthem.

ACKNOWLEDGMENT

The research was supported by funding from the ScottishGovernment Rural and Environment Science and Analyt-ical Services Division (RESAS).

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