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Effects of Clonal Integration on Microbial Community Composition and Processes in the Rhizosphere of the Stoloniferous Herb Glechoma longituba (Nakai) Kuprian

Effects of Clonal Integration on Microbial Community Composition and Processes in the Rhizosphere of the Stoloniferous Herb Glechoma longituba (Nakai) Kuprian

  • Ningfei Lei, 
  • Jun Li, 
  • Shijun Ni, 
  • Jinsong Chen
PLOS
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Abstract

The effects of rhizodeposition on soil C and N availabilities lead to substantial changes of microbial community composition and processes in the rhizosphere of plants. Under heterogeneous light, photosynthates can be translocated or shared between exposed and shaded ramets by clonal integration. Clonal integration may enhance the rhizodeposition of the shaded ramets, which further influences nutrient recycling in their rhizosphere. To test the hypothesis, we conducted a pot experiment by the stoloniferous herb Glechoma longituba subjected to heterogeneous light. Microbial biomass and community composition in the rhizosphere of shaded offspring ramets, assessed by phospholipid fatty acids (PLFAs) analysis, were markedly altered by clonal integration. Clonal integration positively affected C, N availabilities, invertase and urease activities, N mineralization (Nmin) and nitrification rates (Nnitri) in the rhizosphere of shaded offspring ramets. However, an opposite pattern was also observed in phenoloxidase (POXase) and peroxidase (PODase) activities. Our results demonstrated that clonal integration facilitated N assimilation and uptake in the rhizosphere of shaded offspring ramets. The experiment provides insights into the mechanism of nutrient recycling mediated by clonal integration.

Introduction

Rhizosphere, a zone of usually high microbial turnover and activity, has been coined to describe the soil adjacent to and influenced by plant roots [1]. Plant-derived root exudates are primary sources of labile C inputting to soil [2], [3]. These labile C sources rapidly metabolized by microorganisms may generally stimulate their growth or succession in the rhizosphere [4], [5]. So, plant roots exert strong influences on the rhizosphere through ‘rhizodeposition’ (root exudation such as sugars, amino acids, organic acids and hormones, as well as mucilage, enzymes, sloughed root cells and C allocated to root-associated symbionts) [6].

In the form of rhizodeposition, photosynthates released into soil by plant roots are a major source of carbon, energy or structural material for soil microorganisms and affect the microbial community composition in the rhizosphere [7][11]. Fungi, especially ectomycorrhizal (ECM) fungi involved in nitrogen turnover (e.g. mineralization and nitrification), prefer the substrates with larger C/N ratios [12]. Microbial processes, such as extracellular enzyme activities [1], [13], [14], N mineralization and nitrification [15], [16], are mediated by specific groups of microorganisms in the rhizosphere. Two experiments to disrupt root exudation into the soil demonstrated that decreased resource availability negatively affected nitrogen mineralization and nitrification in the rhizosphere via rhizodeposition from plant root [1], [15]. So, microbial processes are highly sensitive to the availabilities of labile C and N in the rhizosphere [5].

Clonal plants can translocate or share resources, such as carbohydrates, water and nutrients among interconnected ramets through clonal integration [17]. Shading may have negative effects on photosynthetic capacity and growth performance of plants [18], [19]. Clonal integration may alter resource levels of ramets under heterogeneous habitats [20], [21]. So, enhanced photosynthates availability caused by clonal integration may have a significant influence on microbial community composition and processes in the rhizosphere of the ramets subjected to low light availability stress. Further, microorganisms present in the rhizosphere may mediate nutrient availability for plants by carrying out a wide spectrum of decomposition processes. As mentioned-above, rhizosphere processes may play a vital role in community or ecosystem nutrient cycling [1]. However, studies on the mechanism of nutrient recycling mediated by clonal integration are rare in the rhizosphere. A pot experiment was conducted by the stoloniferous herb Glechoma longituba subjected to heterogeneous light (mother ramets suffering from full sun versus offspring ones suffering from 80% shade). Comparing with severed offspring ramets, we predicted that connected offspring ramets displayed (1) higher C and N availabilities in the rhizosphere. Based on effects of C and N availabilities on microbial community composition and processes, we expected that connected offspring ramets exhibited (2) higher microbial biomass and different microbial community composition in the rhizosphere; (3) higher extracellular enzymes activities in the rhizosphere; (4) greater N mineralization and nitrification rates in the rhizosphere.

Material and Methods

Plant species and experimental design

G. longituba (Lamiaceae) is a stoloniferous perennial herb. Its monopodial stolons are able to creep on the ground. Ramets can develop on all stolon nodes. A genet or fragment consists of a number of ramets connected by stolons for a certain period of time. Each ramet has two zygomorphic single leaves originating from a stolon node. Every leaf axil bears one bud that may grow into a secondary stolon. The plant is generally found in forests, on roadsides or by creeks and distributed all over China except for the Northwest [22].

In May 2012, ten original clonal fragments of G. longituba were collected from a forest understory in Suining City (30°10′∼31°10′N; 105°03′∼106°59′E), Sichuan Province, China. The sampling site did not belong to the part of any farms or national parks. G. longituba is widespread in China and not an endangered or protected species, so we did not need any relevant permissions/permits for plant sample collection. These original plants were at least 100 m apart from one another. They were propagated in a greenhouse with a mean temperature of 22±8°C. The plants were watered and fertilized as needed.

In June 2013, each clonal fragment consisting of a mother and an offspring ramet with similar size was selected. The two ramets were planted separately in two adjacent plastic pots (10 cm in diameter, 8.5 cm in height) filled with a 3∶1 mixture of humus soil and sand. Plants were watered regularly with distilled water to prevent water stress. After two weeks of growth, offspring ramets were subjected to a 80% shading treatment and the other mother ramets were grown in full sun, whereas stolons between the mother and the offspring ramets were either severed or remained intact (Fig. 1). No new offspring ramets were produced during the two weeks of recovery. Shading was imposed by placing small shade cages covered with black cloth above the pots. The mesh was covered on the top of each pot to avoid potential effects of litters. Only original ramets were allowed to root during the experiment. Each treatment was replicated 10 times and all treatments included clonal fragments from the 10 original plants. All replicates of each treatment were randomly located on benches in a greenhouse. Because soil closely adhering to the roots (up to 2 mm around the root) was considered as rhizosphere soil, the experimental procedure was repeated 10 times to collect enough soil sample. The experiment lasted for 10 weeks.

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Figure 1. Schematic diagram of the experimental design.

The clonal fragment consists of mother ramet and its successional offspring ramet. The offspring ramets were subjected to the 80% shading treatment, and the mother ramets were exposed to full sun. The stolon between mother and offspring ramets remained intact or severed. The mesh was covered on the top of each pot to avoid potential effects of litters.

https://doi.org/10.1371/journal.pone.0108259.g001

Soil sampling and assays

At the end of experiment, rhizosphere soil was sampled according to the shaking root method [23]. The rhizosphere soil of shaded offspring ramets was separated from roots by hand, sieved (<2 mm mesh) and stored at −20°C.

Soil microbial biomass carbon (Cmic) and microbial biomass nitrogen (Nmic) were analyzed using chloroform fumigation-extraction (CFE) method [24], [25]. Briefly, 20 g of fresh, sieved soil was used for the fumigation and non-fumigation treatments, both extracted using 0.5 M K2SO4 at a ratio of 1∶4 (w/v), shaken for 30 min and filtrated through a Whatman no. 42 filter paper. The K2SO4-extract of both fumigated and non-fumigated samples were analyzed immediately for dissolved organic carbon (DOC) and dissolved organic nitrogen (DON) using a TOC/TN analyzer (elementar vario TOC SELECT, Germany). Cmic and Nmic were calculated using the following equations: Cmic (or Nmic) = 2.22×EB [26], where EB was the difference of carbon (or nitrogen) extracted from fumigated soil between non-fumigated soil. Soil moisture was detected gravimetrically, i.e. a sample of 20 g was oven-dried at 105°Cfor 48 h until a constant weight. Total soil organic carbon (TOC) and total nitrogen (TN) were determined using an elemental analyzer (elementar vario MACRO CUBE, Germany). Soil pH was measured in a ratio of 1: 2.5 (soil: water, w/v).

N mineralization and nitrification

N mineralization and nitrification were assessed by the modified anaerobic incubation method [27]. Briefly, fresh soil samples (5 g) were placed into a 200 mL plastic bottles, 10 mL deionized water was added to the bottles to thoroughly submerge the soil. The plastic bottles were sealed with stopper to avoid water evaporation during incubation, and placed in a constant temperature (40°C) incubator for 7 days. At the beginning of the incubation experiment, pre-incubation soil was sampled to measure the initial concentrations of NH4+-N and NO3-N. After a week of incubation, the post-incubation soil samples were mixed with 40 mL of 2 M KCl using a 1: 8 soil: extractant (w/v) ratio, shaken for 30 min on a reciprocal shaker; then the extracts were filtered through prewashed Whatman no. 42 filter papers and supernatants were stored at −20°C until analysis of NH4+-N and NO3-N concentrations. The NH4+-N and NO3-N concentrations were separately measured by spectrophotometry using the ammonium indophenol blue method and the cadmium reduction method [28]. All concentrations of NH4+-N and NO3-N were based on dry soil weight and expressed on a mg·g−1 (DW). The N mineralization rate (Nmin) was calculated as the changes in the inorganic N (NH4+-N, NO3-N) content from time zero to 7 days. A similar formula was used to calculate N nitrification rate (Nnitri) [29], [30]:

Extracellular enzymes activities assays

Invertase activity was measured by the modified method [31]. Briefly, fresh soil (1.0 g) was added to 5 mL 0.1 M saccharose in 0.1 M Na-acetate buffer (pH 4.65) in a 50 mL reaction flask and incubated for 1 h at 30°C. After of incubation, the mixture was transferred immediately to a freezer for 10 min to stop the enzymatic reaction, centrifuged at 3500 g for 10 min and the reaction products were determined in the supernatants. The concentrations of glucose produced by saccharose hydrolysis were determined by the Nelson-Somoji reagent at 660 nm [32]. Invertase activity was expressed in µg glucose g−1DW h−1.

For the determination of urease activity, the modified procedure was adopted according to the description [33]. Briefly, 5 g fresh soil was incubated with 2.5 mL 0.08 M urea solution and 20 mL borate buffer (pH 10.0) for 2 h at 37°C. Released ammonium was extracted using 50 mL 2 M KCl solution, and determined colorimetrically at 690 nm.

POXase activity was measured according to an improved procedure [34]. Briefly, 1.0 g of fresh soil was added to 3 mL of reagent solution (obtained by mixing 1.5 mL of catechol solution with 1.5 mL of proline solution) and 2 mL of phosphate buffer (0.1 M, pH 6.5). The suspension was swirled and incubated at 37°C for 1 h, then reaction was stopped by cooling and adding 5 mL of ethanol. The mixture was centrifuged at 4000 g at 4°C for 5 min. The absorbance of the supernatant fraction was determined at 525 nm. Assays without soil and catechol were carried out simultaneously as controls. POXase activity was expressed as µmol oxidized catechol (o-catechol) g−1DW h−1.

PODase activity was determined with 3,3′,5,5′-tetramethylbenzidine (TMB) as the substrates [35]. Samples of fresh soil (4 g) were mixed with 200 mL cold acetate buffer (5°C, 50 mM, pH 5.0) on a vortex mixer for 30 s at high speed. The soil suspension was then diluted 20-fold in acetate buffer. Aliquots of 0.25 mL were transferred to 2 mL centrifuge tubes. 0.5 mL preheated TMB reagent (25°C) was added to the centrifuge tubes and the tubes were incubated in a constant temperature (25°C) incubator. After 2 h, peroxidase reaction was terminated by adding 1.2 mL sulfuric acid (0.2 M). The tubes were then centrifuged under dark and absorbance of the supernatants were read at 450 nm. Controls were performed using acetate buffer substituted TMB to confirm that there was no photo-oxidation of TMB.

Microbial community composition

Microbial community composition was assessed using phospholipid fatty acids (PLFAs) [1]. (1) Extraction: A sample of 8.0 g fresh soil was extracted with a mixture of citrate buffer (0.15 M, pH 4.0 with NaOH), chloroform and methanol at a ratio of 0.8: 1: 2 (v/v/v). Suspension was shaken darkly at 25°C for 30 min, centrifuged at 10000 g for 0.5 h and the supernatant was transferred to new vials. Chloroform and citrate buffer were added to supernatant for separation of the phases. After 18 h, the organic phases were removed and dried under a stream of dry N2. (2) Chromatography: lipids were redissolved in chloroform and neutral lipids were separated from phospholipids on silica columns by elution with chloroform (5 mL), acetone (10 mL) and methanol (5 mL) gradually. Methanol-phase was collected and dried under N2 stream. (3) Methyl esterification: phospholipids were subsequently converted to fatty acids methyl esters (FAMEs) by alkaline methanolysis. Phospholipids were dissolved in 1 mL of methanolic 0.2 M KOH and 1 mL of methanol-toluene (1∶1, v/v) and incubated for 15 min at 37°C, then mixed with 2 mL of deionized water, 0.3 mL of acetic acids (0.2 M) and 2 mL of hexane, swirled and centrifuged for 10 min. The hexane-phase was removed and dried under N2 stream. After adding 100 µL of a solution of methyl-nonadecanoate (C19:0, 25 ng µL−1) as an internal standard, FAMEs were dissolved in C19:0 and analyzed by capillary gas chromatography (Agilent Technologies, 6850N-GC System, USA). Concentration of single FAMEs was calculated using the internal standard (C19:0) peak as a reference according to the following formula:

Where A and B were the peak areas of each fatty acid methyl ester and internal standard, respectively; W was the oven-dry soil weight (DW).

PLFAs used as biomarkers for specific groups of soil microorganisms and were designated according to an standard nomenclature: ((a,i,cy)X: YωZ(c,t)), where the X referred to the number of C atoms, the Y indicated the number of double bonds followed by the position (ω) and distance (Z) of the double bonds from the methyl end. The prefixes a, i indicated anteiso- and iso-branching; the suffixes c, t referred to cis- and trans-double bonds; cy represented cyclopropyl-group. The fatty acids used as biomarkers for specific groups of soil organisms were listed in Table 1.

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Table 1. PLFAs biomarkers of different microbial groups were used in the study.

https://doi.org/10.1371/journal.pone.0108259.t001

Statistical analysis

Soil pH, moisture, C and N availabilities, microbial biomass, extracellular enzymes activities, N mineralization and nitrification rates in the rhizosphere of shaded offspring ramets were investigated by one-way ANOVA. Microbial community composition of shaded offspring ramets was analyzed by a principal component analysis (PCA) using specific PLFAs biomarkers. For PCA analysis, scores of different soil microbial groups were expressed as percentage of the total PLFAs in the sample. Pearson correlations were used for relating PLFAs concentrations of different microbial groups to extracellular enzymes activities, N mineralization and nitrification, C and N availabilities. Significance was set at p = 0.05 level. If needed, data were natural logarithm-transformed or arcsine-transformed in order to achieve normality and homogeneity of variance. All statistical analyses were performed using SPSS 20.0 software (SPSS, Chicago, IL, USA).

Results

Changes in soil properties

Clonal integration significantly increased TOC, DOC and DON concentrations in the rhizosphere of shaded offspring ramets as well as C/N, whereas no effects of clonal integration on soil moisture, pH and TN were observed in the rhizosphere of shaded offspring ramets (Table 2). Cmic and Nmic were significantly higher in the rhizosphere of shaded, connected offspring ramets compared to shaded, severed offspring ramets (Table 2). Concentrations of inorganic nitrogen (NH4+-N and NO3-N) in the rhizosphere of shaded offspring ramets were also markedly increased by clonal integration. Meanwhile, NH4+-N concentration was evidently higher than NO3-N concentration, regardless of stolon connection or severing (Table 2).

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Table 2. Effects of clonal integration on soil properties in the rhizosphere of shaded offspring ramets.

https://doi.org/10.1371/journal.pone.0108259.t002

Changes in microbial community composition

A principal component analysis (PCA) based on PLFAs biomarkers of different microbial groups revealed that microbial community composition was clearly distinct in the rhizosphere of shaded, connected offspring ramets compared to shaded, severed offspring ramets (Fig. 2). The results of PCA were further supported by the absolute PLFAs concentrations for different microbial groups (Fig. 3). The PLFAs concentrations of bacteria, fungi, Gram-positive bacteria and Gram-negative bacteria were significantly increased by clonal integration as well as the total PLFAs concentration (Fig. 3a,b,c,e,f). The biomarker (18:2ω6,9) accounted for 86% of total fungal PLFAs concentrations in the rhizosphere of shaded, connected offspring ramets, the proportion was slightly decreased (75%) in shaded, severed offspring ones. Specially, clonal integration distinctly increased the PLFAs concentration of ECM fungi (18:2ω6,9) by 72% (Fig. 3g). However, effects of clonal integration on the PLFAs concentration of actinomycete and Ba/Fu were not observed (Fig. 3d, h).

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Figure 2. Microbial community composition described by a principal component analysis of the concentrations of PLFAs.

Error bars showed standard error of the mean of PCA weighted loading values for connected (C) and severed (S) shaded offspring ramets. Black squares represented PCA weighted loading values of microorganisms. Microbial groups abbreviations: TP, total PLFAs; Ba, bacteria; Fu, fungi; Ac, actinomycete; G+, gram-positive bacteria; G, gram-negative bacteria, n = 10.

https://doi.org/10.1371/journal.pone.0108259.g002

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Figure 3. Concentrations or ratios of soil microbial groups PLFAs in the rhizosphere of shaded offspring ramets.

Significant differences between connected offspring ramets (black bars) and severed offspring ones (open bars) were indicated by *** (p<0.001), ** (p<0.01), * (p<0.05) and ns (not significant). Error bars represented standard errors, n = 10.

https://doi.org/10.1371/journal.pone.0108259.g003

Changes in microbial processes

Compared to shaded, severed offspring ramets, invertase and urease activities in the rhizosphere of shaded, connected offspring ramets were increased by 59.8% (p<0.001) and 38.9% (p = 0.005) respectively (Fig. 4a,b). On the contrary, POXase and PODase activities in the rhizosphere of shaded, connected offspring ramets were decreased by 47.8% (p<0.001) and 12.7% (p<0.001) respectively (Fig. 4c,d). Nnitri accounted for only 17% of Nmin in the rhizosphere of shaded, connected offspring ramets, compared to 28% in the rhizosphere of shaded, severed offspring ones (p = 0.034). Nmin and Nnitri in the rhizosphere of shaded offspring ramets were markedly increased by clonal integration (Fig. 5a,b).

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Figure 4. Extracellular enzymes activities involved in the depolymerization of C, N from SOM.

Invertase (a), Urease (b), POXase (c) and PODase (d) were measured in the rhizosphere of shaded offspring ramets. Significant differences between connected offspring ramets (black bars) and severed offspring ones (open bars) were indicated by *** (p<0.001) and ** (p<0.01). Error bars represented standard errors, n = 10.

https://doi.org/10.1371/journal.pone.0108259.g004

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Figure 5. N mineralization rate (a) and nitrification rate (b) in the rhizosphere of shaded offspring ramets were measured by anaerobic incubation methods.

Significant differences between connected offspring ramets (black bars) and severed offspring ones (open bars) were indicated by *** (p<0.001), ** (p<0.01). Error bars represented standard errors, n = 10.

https://doi.org/10.1371/journal.pone.0108259.g005

Correlations between the PLFAs concentrations of different microbial groups and soil properties or microbial processes

Soil properties (such as DOC, DON, TOC, NH4+, NO3) were positively correlated to the PLFAs concentrations of most microbial groups except for actinomycete (Table 3). Similarly, microbial processes (such as Nnitri, Nmin, invertase and urease activities) were positively correlated to the PLFAs concentrations of most microbial groups (Table 3). On the contrary, POXase and PODase activities were negatively correlated to the PLFAs concentrations of most microbial groups (Table 3). In addition, Ba/Fu and G/G+ ratios were not significantly correlated with any of the measured soil properties and microbial processes (Table 3).

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Table 3. Correlations between concentrations or ratios of soil microbial groups PLFAs and soil properties or microbial processes in the rhizosphere of shaded offspring ramets.

https://doi.org/10.1371/journal.pone.0108259.t003

Discussion

Changes in C and N availabilities

A major source of labile C inputting to soil is the root exudates [3], [36], [37]. Because girdling blocked the flow of newly formed photosynthates to the roots and to mycorrhizal fungi, it significantly decreased concentration of DOC and Cmic in the rhizosphere or bulk soils [1], [13], [38]. Clonal plants can translocate or share photosynthates from exposed ramets to shaded ramets by clonal integration [39]. So, effects of clonal integration on labile C may be similar with those reported in previous girdling experiments. Root exudation is associated with increased N availability in the rhizosphere [14]. DON and Nmic were decreased in the girdled plots [38]. The decrease in DOC and DON in girdled plots was probably caused by the decrease in root exudation because a substantial portion of DOC and DON originated from photosynthates [14]. In our study, the increase of belowground carbon allocation caused by clonal integration may improve C and N availability in the rhizosphere of shaded offspring ramets.

Increased dissolved inorganic N concentrations in response to girdling have been found in other studies [14], [40]. On the contrary, we suspect that clonal integration, by increasing the supply of available C to microorganisms, may stimulate N mineralization and nitrificantion in the rhizosphere of shaded offspring ramets. Alternatively, effects of available C supply on soil dissolved inorganic N concentrations may depend on the species-specific. Compared with NH4+-N, NO3-N concentration was lower in the rhizosphere of shaded offspring ramets, regardless of stolon connection or severing (Table 2). This is most likely due to the high mobility of NO3 in soil [40].

Changes in microbial biomass and community composition

DOC is an available C source for microbes and affects their abundance, composition and activity [41]. A larger portion of easily assimilable C was derived from photosynthates produced by plants and shaped a specific microbial community composition [9], [11], [42],[43]. Tree girdling did not affect the total PLFAs concentration and increased the concentration of bacterial PLFAs [38]. In addition, the concentrations of PLFAs for bacterial groups were also related to soil pH [44]. Although no effect of clonal integration on soil pH was observed, the total, bacteria, Gram-positive bacteria and Gram-negative bacteria PLFAs concentrations in the rhizosphere of shaded offspring ramets were significant increased by clonal integration (Table 2; Fig. 3). The similar patterns were observed in another girdling experiment [1].

Specially, clonal integration distinctly increased the PLFAs concentration of ECM fungi (18:2ω6,9) by 72% (Fig. 3g). This was consistent with a previous girdling experiment [13]. Our results further confirm that the dramatic increase in fungi caused by clonal integration is associated with a increase in C supply and that fungi depend to a much higher degree on belowground C allocation [12].

The increase of total fungal biomass caused by clonal integration was mainly related to the increase of ECM fungal biomass [38], [41]. Compared to Gram-positive bacterial PLFAs, the concentrations of Gram-negative bacterial PLFAs were higher in the rhizosphere of shaded offspring ramets, regardless of stolon connection or severing (Fig. 3e,f). This is consistent with the suggestion that Gram-negative bacteria are generally favored by the labile C substrates released by rhizosdeposition and more frequent in the rhizosphere [1]. The effects of clonal integration on actinomycetes were not observed (Fig. 3d). A possible explanation is that actinomycetes are less stimulated in the rhizosphere [45]. Association relationships between different microbial groups (bacteria, Gram-negative bacteria, Gram-positive bacteria, fungi and ECM) and DOC or TOC concentrations may imply their strong dependence (Table 3). We tentatively conclude that the effects of clonal integration on soil C and N availability lead to substantial changes in microbial biomass and community composition in the rhizosphere of shaded offspring ramets.

Changes in microbial processes

Soil enzyme activities can be used as potential indicators of nutrient cycling processes. Invertase catalyzes hydrolytic processes of SOM [44]. Urease is generally produced by bacteria, filamentous fungi and yeasts, thus enhancing N mineralization [46], [47]. Plant root exudates may provide a constant energy supply, thereby creating optimal conditions for SOM degraders [48]. The increased microbial biomass caused by enhancing root exudation could increase extracellular enzymes activities and the release of N from SOM [49], [50]. The suggestions were further supported by the positive correlations between invertase or urease activities and most soil microbial groups in the rhizosphere of shaded offspring ramets (Table 3).

Phenoloxidase (POXase) and peroxidase (PODase) are the lignolytic enzyme involved in the degradation of recalcitrant SOM (e.g. lignin) [51]. POXase and PODase are generally produced by slow-growing specialist decomposers (e.g. saprotrophic fungi) [52], [53]. Competition between microbial groups could also have been responsible for the shift of enzyme activities. Mycorrhizal fungi are known to dominate the rooted soil layers as a result of a competitive advantage gained through access to root C, whereas saprotrophic fungi are thought to be more competitive in the litter layer [53], [54]. Clonal integration greatly increased the abundance of mycorrhizal fungi, thereby possibly also giving saprotrophic fungi a competitive disadvantage in the rooting zone. The suggestions were further confirmed by the negative correlations between POXase or PODase activities and most soil microbial groups in the rhizosphere of shaded offspring ramets (Table 3). The similar patterns were observed in a previous girdling experiment [1].

Dissolved organic matter (e.g. DOC, DON) was considered to influence soil microbial processes, such as soil respiration/C mineralization [55], [56] and N mineralization [57], [58]. N mineralization and nitrification were regulated by a variety of heterotrophic bacteria and fungi via using labile C source or SOM [40]. N mineralization and nitrification rates were strongly increased by clonal integration in the rhizosphere of shaded offspring ramets (Fig. 5). Positive correlations between N mineralization or nitrification rates and most microbial groups were observed in the rhizosphere of shaded offspring ramets (Table 3). The similar patterns were found in a previous girdling experiment [1]. Notably, Nnitri tended to be much lower than Nmin in the rhizosphere of shaded offspring ramets, regardless of stolon connection or severing (Fig. 5). This could be explained by the fact that NO3 was derived from the oxidizing of NH4+ (i.e. nitrification) by chemoautotrophic bacteria or heterotrophic microorganisms [59], [60]. In addition, NO3 was water soluble and seldom present in detectable amounts [61].

Because of a high availability of easily assimilable carbon and nutrients, microbial community composition were modified by clonal integration in the rhizosphere of shaded offspring ramets. Invertase and urease activitties, N mineralization and nitrification rates were enhanced by clonal integration in the rhizosphere of shaded offspring ramets. So, clonal integration may facilitate N assimilation and uptake in the rhizosphere of shaded offspring ramets. A field study investigated the effects of clonal integration on nutrient recycling of the Serengeti grassland communities [62]. Our experiment provides insights into the mechanism of nutrient recycling mediated by clonal integration. To allow a robust generalization, however, more experimental studies, especially those conducted in the field, are required.

Author Contributions

Conceived and designed the experiments: NFL JL JSC. Performed the experiments: NFL JL. Analyzed the data: SJN JSC. Contributed reagents/materials/analysis tools: NFL JL. Wrote the paper: NFL JL. JL SJN JSC.

References

  1. 1. Koranda M, Schnecker J, Kaiser C, Fuchslueger L, Kitzler B, et al. (2011) Microbial processes and community composition in the rhizosphere of European beech - The influence of plant C exudates. Soil Biology and Biochemistry 43: 551–558.
  2. 2. Hütsch BW, Augustin J, Merbach W (2002) Plant rhizodeposition—an important source for carbon turnover in soils. Journal of Plant Nutrition and Soil Science 165: 397–407.
  3. 3. Kuzyakov Y (2002) Review: Factors affecting rhizosphere priming effects. Journal of Plant Nutrition and Soil Science 165: 382–396.
  4. 4. Butler JL, Bottomley PJ, Griffith SM, Myrold DD (2004) Distribution and turnover of recently fixed photosynthate in ryegrass rhizospheres. Soil Biology and Biochemistry 36: 371–382.
  5. 5. Schimel JP, Weintraub MN (2003) The implications of exoenzyme activity on microbial carbon and nitrogen limitation in soil: a theoretical model. Soil Biology and Biochemistry 35: 549–563.
  6. 6. Bais HP, Weir TL, Perry LG, Gilroy S, Vivanco JM (2006) The role of root exudates in rhizosphere interactions with plants and other organisms. Annual Review of Plant Biology 57: 233–266.
  7. 7. Barea J-M, Pozo MJ, Azcon R, Azcon-Aguilar C (2005) Microbial cooperation in the rhizosphere. Journal of Experimental Botany 56: 1761–1778.
  8. 8. Fu S, Cheng W (2004) Defoliation affects rhizosphere respiration and rhizosphere priming effect on decomposition of soil organic matter under a sunflower species: Helianthus annuus. Plant and Soil 263: 345–352.
  9. 9. Grayston S, Vaughan D, Jones D (1997) Rhizosphere carbon flow in trees, in comparison with annual plants: the importance of root exudation and its impact on microbial activity and nutrient availability. Applied Soil Ecology 5: 29–56.
  10. 10. Kuzyakov Y, Cheng W (2001) Photosynthesis controls of rhizosphere respiration and organic matter decomposition. Soil Biology and Biochemistry 33: 1915–1925.
  11. 11. Rajaniemi T, Allison V (2009) Abiotic conditions and plant cover differentially affect microbial biomass and community composition on dune gradients. Soil Biology and Biochemistry 41: 102–109.
  12. 12. Olsson PA (1999) Signature fatty acids provide tools for determination of the distribution and interactions of mycorrhizal fungi in soil. FEMS Microbiology Ecology 29: 303–310.
  13. 13. Kaiser C, Koranda M, Kitzler B, Fuchslueger L, Schnecker J, et al. (2010) Belowground carbon allocation by trees drives seasonal patterns of extracellular enzyme activities by altering microbial community composition in a beech forest soil. New Phytologist 187: 843–858.
  14. 14. Weintraub MN, Scott-Denton LE, Schmidt SK, Monson RK (2007) The effects of tree rhizodeposition on soil exoenzyme activity, dissolved organic carbon, and nutrient availability in a subalpine forest ecosystem. Oecologia 154: 327–338.
  15. 15. Norton JM, Firestone MK (1996) N dynamics in the rhizosphere of Pinus ponderosa seedlings. Soil Biology and Biochemistry 28: 351–362.
  16. 16. Priha O, Grayston SJ, Pennanen T, Smolander A (1999) Microbial activities related to C and N cycling and microbial community structure in the rhizospheres of Pinus sylvestris, Picea abies and Betula pendula seedlings in an organic and mineral soil. FEMS Microbiology Ecology 30: 187–199.
  17. 17. Du J, Wang N, Alpert P, Yu MJ, Yu FH, et al. (2010) Clonal integration increases performance of ramets of the fern Diplopterygium glaucum in an evergreen forest in southeastern China. Flora 205: 399–403.
  18. 18. Roiloa SR, Retuerto R (2007) Responses of the clonal Fragaria vesca to microtopographic heterogeneity under different water and light conditions. Environmental and Experimental Botany 61: 1–9.
  19. 19. He WM, Alpert P, Yu FH, Zhang LL, Dong M (2011) Reciprocal and coincident patchiness of multiple resources differentially affect benefits of clonal integration in two perennial plants. Journal of Ecology 99: 1202–1210.
  20. 20. Alpert P, Mooney H (1986) Resource sharing among ramets in the clonal herb, Fragaria chiloensis. Oecologia 70: 227–233.
  21. 21. Evans JP (1991) The effect of resource integration on fitness related traits in a clonal dune perennial, Hydrocotyle bonariensis. Oecologia 86: 268–275.
  22. 22. Liao M, Yu F, Song M, Zhang S, Zhang J, et al. (2003) Plasticity in R/S ratio, morphology and fitness-related traits in response to reciprocal patchiness of light and nutrients in the stoloniferous herb, Glechoma longituba L. Acta Oecologica 24: 231–239.
  23. 23. Riley D, Barber S (1970) Salt accumulation at the soybean (Glycine max (L.) Merr.) root-soil interface. Soil Science Society of America Journal 34: 154–155.
  24. 24. Vance E, Brookes P, Jenkinson D (1987) An extraction method for measuring soil microbial biomass C. Soil biology and Biochemistry 19: 703–707.
  25. 25. Brookes P, Landman A, Pruden G, Jenkinson D (1985) Chloroform fumigation and the release of soil nitrogen: a rapid direct extraction method to measure microbial biomass nitrogen in soil. Soil Biology and Biochemistry 17: 837–842.
  26. 26. Wu J, Joergensen R, Pommerening B, Chaussod R, Brookes P (1990) Measurement of soil microbial biomass C by fumigation extractionan - automated procedure. Soil Biology and Biochemistry 22: 1167–1169.
  27. 27. Carter MR, Gregorich EG (1993) Soil sampling and methods of analysis. Canadian: Canadian Society of Soil Science. 599–605 p.
  28. 28. Carter MR, Gregorich EG (1993) Soil sampling and methods of analysis. Canadian: Canadian Society of Soil Science. 71–80 p.
  29. 29. Zhou W, Chen H, Zhou L, Lewis BJ, Ye Y, et al. (2011) Effect of freezing-thawing on nitrogen mineralization in vegetation soils of four landscape zones of Changbai Mountain. Annals of Forest Science 68: 943–951.
  30. 30. Ren W, Chen F-s, Hu X-f, Yu M-q, Feng X (2011) Soil nitrogen transformations varied with plant community under Nanchang urban forests in mid-subtropical zone of China. Journal of Forestry Research 22: 569–576.
  31. 31. Gianfreda L, Sannino F, Violante A (1995) Pesticide effects on the activity of free, immobilized and soil invertase. Soil Biology and Biochemistry 27: 1201–1208.
  32. 32. Nelson N (1944) A photometric adaptation of the Somogyi method for the determination of glucose. Journal of Biological Chemistry 153: 375–379.
  33. 33. Kandeler E, Gerber H (1988) Short-term assay of soil urease activity using colorimetric determination of ammonium. Biology and Fertility of Soils 6: 68–72.
  34. 34. Perucci P, Casucci C, Dumontet S (2000) An improved method to evaluate the o-diphenol oxidase activity of soil. Soil Biology and Biochemistry 32: 1927–1933.
  35. 35. Johnsen AR, Jacobsen OS (2008) A quick and sensitive method for the quantification of peroxidase activity of organic surface soil from forests. Soil Biology and Biochemistry 40: 814–821.
  36. 36. Jones DL, Hodge A, Kuzyakov Y (2004) Plant and mycorrhizal regulation of rhizodeposition. New Phytologist 163: 459–480.
  37. 37. Jones D, Nguyen C, Finlay R (2009) Carbon flow in the rhizosphere: carbon trading at the soil–root interface. Plant and Soil 321: 5–33.
  38. 38. Chen D, Zhou L, Wu J, Hsu J, Lin Y, et al. (2012) Tree girdling affects the soil microbial community by modifying resource availability in two subtropical plantations. Applied Soil Ecology 53: 108–115.
  39. 39. Alpert P ((1999)) Effects of clonal integration on plant plasticity in Fragaria chiloensis. Plant Ecology 141: 99–106.
  40. 40. Zeller B, Liu J, Buchmann N, Richter A (2008) Tree girdling increases soil N mineralisation in two spruce stands. Soil Biology and Biochemistry 40: 1155–1166.
  41. 41. Högberg MN, Högberg P, Myrold DD (2007) Is microbial community composition in boreal forest soils determined by pH, C-to-N ratio, the trees, or all three? Oecologia 150: 590–601.
  42. 42. Brant JB, Myrold DD, Sulzman EW (2006) Root controls on soil microbial community structure in forest soils. Oecologia 148: 650–659.
  43. 43. Yarwood SA, Myrold DD, Högberg MN (2009) Termination of belowground C allocation by trees alters soil fungal and bacterial communities in a boreal forest. FEMS Microbiology Ecology 70: 151–162.
  44. 44. Gu Y, Wang P, Kong CH (2009) Urease, invertase, dehydrogenase and polyphenoloxidase activities in paddy soil influenced by allelopathic rice variety. European Journal of Soil Biology 45: 436–441.
  45. 45. Zhang LZ, Niu W, Niu Y, Niu XW ((2009)) Impact of Caragana Fabr. plantation on plant community and soil properities of saline-alkali wasteland. Acta Ecologica Sinica 29(9): 4693–4699.
  46. 46. Mobley H, Hausinger R (1989) Microbial ureases: significance, regulation, and molecular characterization. Microbiological Reviews 53: 85–108.
  47. 47. Gianfreda L, Antonietta Rao M, Piotrowska A, Palumbo G, Colombo C (2005) Soil enzyme activities as affected by anthropogenic alterations: intensive agricultural practices and organic pollution. Science of the Total Environment 341: 265–279.
  48. 48. Fontaine S, Mariotti A, Abbadie L (2003) The priming effect of organic matter: a question of microbial competition? Soil Biology and Biochemistry 35: 837–843.
  49. 49. Cheng W, Kuzyakov Y (2005) Root effects on soil organic matter decomposition. American society of Agronomy 48: 119–143.
  50. 50. Blagodatskaya E, Blagodatsky S, Dorodnikov M, Kuzyakov Y (2010) Elevated atmospheric CO2 increases microbial growth rates in soil: results of three CO2 enrichment experiments. Global Change Biology 16: 836–848.
  51. 51. Sinsabaugh RL (2010) Phenol oxidase, peroxidase and organic matter dynamics of soil. Soil Biology and Biochemistry 42: 391–404.
  52. 52. Baldrian P (2009) Ectomycorrhizal fungi and their enzymes in soils: is there enough evidence for their role as facultative soil saprotrophs? Oecologia 161: 657–660.
  53. 53. Hobbie EA, Horton TR (2007) Evidence that saprotrophic fungi mobilise carbon and mycorrhizal fungi mobilise nitrogen during litter decomposition. New Phytologist 173: 447–449.
  54. 54. Lindahl BD, Ihrmark K, Boberg J, Trumbore SE, Högberg P, et al. (2007) Spatial separation of litter decomposition and mycorrhizal nitrogen uptake in a boreal forest. New Phytologist 173: 611–620.
  55. 55. Chantigny MH, Angers DA, Prévost D, Simard RR, Chalifour F-P (1999) Dynamics of soluble organic C and C mineralization in cultivated soils with varying N fertilization. Soil Biology and Biochemistry 31: 543–550.
  56. 56. Gregorich E, Rochette P, McGuire S, Liang B, Lessard R (1998) Soluble organic carbon and carbon dioxide fluxes in maize fields receiving spring-applied manure. Journal of Environmental Quality 27: 209–214.
  57. 57. Appel T, Mengel K (1993) Nitrogen fractions in sandy soils in relation to plant nitrogen uptake and organic matter incorporation. Soil Biology and Biochemistry 25: 685–691.
  58. 58. Murphy D, Macdonald A, Stockdale Ea, Goulding K, Fortune S, et al. (2000) Soluble organic nitrogen in agricultural soils. Biology and Fertility of Soils 30: 374–387.
  59. 59. De Boer W, Kowalchuk G (2001) Nitrification in acid soils: micro-organisms and mechanisms. Soil Biology and Biochemistry 33: 853–866.
  60. 60. Brierley E, Wood M, Shaw P (2001) Influence of tree species and ground vegetation on nitrification in an acid forest soil. Plant and Soil 229: 97–104.
  61. 61. Keeney DR, Nelson D (1982) Methods of soil analysis. Part 2. Chemical and microbiological properties, Nitrogen—inorganic forms. Ameirican: American Society of Agronomy. 643–698.
  62. 62. Wilsey B (2002) Clonal plants in a spatially heterogeneous environment: effects of integration on Serengeti grassland response to defoliation and urine-hits from grazing mammals. Plant Ecology 159: 15–22.
  63. 63. Rinnan R, Michelsen A, Bååth E, Jonasson S (2007) Fifteen years of climate change manipulations alter soil microbial communities in a subarctic heath ecosystem. Global Change Biology 13: 28–39.
  64. 64. Green CT, Scow KM (2000) Analysis of phospholipid fatty acids (PLFA) to characterize microbial communities in aquifers. Hydrogeology Journal 8: 126–141.
  65. 65. Vestal JR, White DC (1989) Lipid analysis in microbial ecology. Bioscience 39: 535–541.