Legume presence reduces the decomposition rate of non-legume roots

Soil Biology & Biochemistry 94 (2016) 88e93

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Legume presence reduces the decomposition rate of non-legume roots Sirgi Saar a, c, Marina Semchenko a, b, Janna M. Barel c, Gerlinde B. De Deyn c, * a

Department of Botany, Institute of Ecology and Earth Sciences, University of Tartu, Lai 40, 51005 Tartu, Estonia Faculty of Life Sciences, University of Manchester, Manchester M13 9PT, United Kingdom c Department of Soil Quality, Wageningen University, P.O. Box 47, 6700 AA Wageningen, The Netherlands b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 22 March 2015 Received in revised form 27 November 2015 Accepted 28 November 2015 Available online 20 December 2015

Living plants can enhance litter decomposition rates via a priming effect by releasing root exudates which provide energy to saprotrophic microbes and thereby enable them to degrade litter faster. The strength of this effect, however, is expected to be dependent on the litter properties. To test whether the presence of a growing plant affects the decomposition rate of dead roots with different traits, we used dead roots of seven species (3 grasses, 3 legumes, 1 forb) as litter and quantified litter mass loss after eight weeks of incubation in soil with or without a growing white clover (Trifolium repens) plant. We expected root decomposition to be faster in the presence of T. repens, especially for roots with high C:N ratio. We found that the presence of T. repens slowed down the decomposition of grass and forb roots (negative priming), while it did not significantly affect the decomposition of legume roots. Our results show that root decomposition can be slowed down in the presence of a living plant and that this effect depends on the properties of the decomposing roots, with a pronounced reduction in root litter poor in N and P, but not in the relatively nutrient-rich legume root litters. Negative priming effect of legume plants on non-legume litter decomposition may have resulted from preferential substrate utilisation by soil microbes. © 2015 Elsevier Ltd. All rights reserved.

Keywords: Litter decomposition Root decomposition Nutrient effects Litter quality Rhizosphere priming effect Plant litter interaction

1. Introduction Litter decomposition and mineralisation are key processes in sustaining primary productivity. Input of litter to soil is derived both from aboveground as well as from belowground plant parts. In temperate grasslands, litter input is larger belowground than aboveground, comprising both root exudates and dead plant roots (Rasse et al., 2005; De Deyn et al., 2008; Freschet et al., 2013). Despite its great relevance, considerably less is known about the decomposition of roots as compared to decomposition of aboveground plant parts. In general, studies of leaf litter decomposition show that environmental conditions (temperature, moisture) and litter quality (concentrations of nutrients, carbon:nitrogen ratio C:N and lignin:N ratios) are most important in determining the speed of litter decomposition (Silver and Miya, 2001; Vivanco and Austin, 2006; Fornara et al., 2009; Freschet et al., 2012a). Leaf litter decomposes fastest in warm and moderately moist conditions and when it is rich in nutrients (i.e. having low C:N ratio) and poor in recalcitrant (i.e. structurally complex) carbon compounds like

* Corresponding author. Tel.: þ31 323174 82123. E-mail address: [email protected] (G.B. De Deyn). http://dx.doi.org/10.1016/j.soilbio.2015.11.026 0038-0717/© 2015 Elsevier Ltd. All rights reserved.

lignin (Taylor et al., 1991). The rate of root litter decomposition and mineralisation can be expected to be driven by the same parameters as leaf litter upon entering the soil. Recent studies focusing on root decomposition found no consistent relationships between root litter decomposition rate and litter N and P concentrations, while a negative relation with lignin concentration and with root C:N was apparent (Silver and Miya, 2001; Freschet et al., 2012a,b; Smith et al., 2014). While direct comparative studies are still scarce, leaf and root decomposition rates have been found to show similar patterns and it seems that chemical composition, including C:N ratio, plays a central role in the process of litter decomposition, irrespective of whether the litter is derived from aboveground or belowground plant parts (Birouste et al., 2012; Freschet et al., 2013). One reason why root decomposition remains poorly studied compared with leaf litter decomposition is that direct quantification of root decomposition as the rate of mass loss represents an inherent challenge as it cannot be performed without disturbing the soil and roots (Silver and Miya, 2001). Most root decomposition studies have used a litter bag approach (Silver and Miya, 2001; Freschet et al., 2012a,b; Smith et al., 2014) which enables comparison of treatment effects between species in a standardised way and gives an indication of potential differential responses though does not mimic in situ decomposition rates. However, this method

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does not reflect in situ decomposition rates as root washing and drying, which is required for accurate estimation of litter mass at the start of decomposition, disrupts the natural rhizosphere community at the soil-root interface. It has been shown that the absolute root decomposition rates are likely to be underestimated when using litterbags and nutrient mobilisation may be affected, especially when working with ectomycorrhizal tree species (Dornbush et al., 2002; Li et al., 2015). However, an alternative method to litter bags such as using intact soil cores remains difficult as the initial mass of roots within cores is unknown and homogenous distribution of roots in the soil is assumed (Dornbush et al., 2002; Li et al., 2015). Living plants can significantly affect the decomposition of soil organic matter and roots through rhizosphere effects, leading to faster or slower decomposition rates (Kuzyakov, 2002, 2010; Fornara et al., 2009; Guenet et al., 2010). The induction of faster decomposition, called positive priming effect, is thought to be governed by root exudates which function as an easily available energy source for soil microbes and enable them to degrade more recalcitrant litter (Kuzyakov, 2002, 2010). Moreover root exudates not only contain C-rich compounds but also N, and both exudate quantity and composition affect microbial activity. Especially root exudates of leguminous plant species tend to be N-rich (Fustec et al., 2011), and can thereby promote growth and activity of soil microorganisms and litter decomposition (Sugiyama and Yazaki, 2012). However, it has also been shown that soil microbes may respond to increased N availability in the soil by reducing the production of enzymes used to mine N from recalcitrant organic matter, thereby slowing down its decomposition (Craine et al., 2007). Also, living plants can slow down litter decomposition, called negative priming (reviewed in Dormaar, 1990), by competing with decomposers for nutrients or by exuding chemical compounds that suppress the decomposers (Van der Krift et al., 2002; Guenet et al., 2010; Kuzyakov, 2010; Coq et al., 2011). Another hypothesis to explain negative priming is the preferential substrate utilization hypothesis (Sparling et al., 1982; Kuzyakov, 2002 for a review). This hypothesis states that the microorganisms actively shift their use of substrates and will use the most easily accessible fresh organic matter as a C source instead of degrading more recalcitrant dead organic matter, which is energetically more costly. To date, the majority of studies have reported positive priming effects of organic matter decomposition by living plants (Cheng et al., 2014) and root exudates have been considered to be a crucial factor in this (Kuzyakov, 2010). However, it is still unclear which traits, both of the living plants and of the litter, cause the wide variation in rhizosphere priming effects. In a large-scale grassland biodiversity experiment, root litter decomposition rate was negatively related to the root biomass but was not affected by the functional group of the plants growing in the soil in which the root litter was decomposing (Fornara et al., 2009). In contrast in another grassland biodiversity experiment, the presence of legumes in the community strongly enhanced the decomposition rate of litter and standard substrates (wooden sticks and cotton strips), indicating positive priming effects on litter decomposition (Scherer-Lorenzen, 2008). These results indicate that the presence of leguminous plant species may generally create positive rhizosphere priming effects of root litter decomposition, but not always, and the underlying mechanisms require further elucidation. Recently it has been proposed that the occurrence of rhizosphere priming effects may strongly depend on whether plants are N- or P-limited, with positive priming effects arising when plants are N-limited and no priming effects when plants are P-limited (Dijkstra et al., 2013). According to this view, one would expect legume species not to have strong priming effects because they are supposedly more limited by P than N as a result of endosymbiosis

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with N2-fixing bacteria. In contrast, Cheng et al. (2003) found that both soybeans (a legume) and spring wheat (a grass) consistently generated positive priming effects, also when ample fertiliser was applied, suggesting that priming effects by plants may not, or not solely, depend on their nutrient status and that also in agricultural systems positive priming effects occur. Root decomposition rates may not only depend on the properties of the root litter or the presence of a living plant, but also on interactive effects between the living plant and the root litter. Van der Krift et al. (2002) showed that the presence of the same focal grass species (Festuca ovina) had differential effects on root litter from different plant species as it accelerated the decomposition of root litter from its own species, but slowed down the decomposition of root litter from another grass species (Anthoxanthum odoratum). The exact underlying mechanisms are still unknown but may be due to different root litter properties that differentially stimulate decomposer organisms. Here we investigated the impact of a growing legume plant on the decomposition rate of different root species and tested the hypotheses that the presence of a growing white clover plant will: 1) increase the decomposition rate of root litter across litter species; 2) the magnitude of the priming effect of the growing plant on root litter decomposition will depend on the initial C:N ratio of the dead roots. We used a litterbag approach to accurately assess decomposition rates. Although being used in most studies of this kind, it has to be noted that overall decomposition rates are likely underestimated and natural rhizosphere communities disturbed compared to in situ situation (Dornbush et al., 2002; Li et al., 2015). 2. Material and methods 2.1. Root litter preparation In preparation of the experiment, seven grassland species comprising three grasses (Lolium perenne, Festuca rubra, Arrhenaterum elatius), three legumes (Trifolium repens, Trifolium pratense, Vicia cracca) and a forb (Cichorium intybus) common in European grasslands were grown on sandy soil at 60% water holding capacity (WHC). The seeds were purchased from specialised companies in The Netherlands: T. pratense and C. intybus from Cruydt-Hoeck (Groningen), L. perenne and T. repens from Agrifirm (Apeldoorn) and in the UK: V. cracca, F. rubra and A. elatius from Emorsegate (Norfolk). The soil was collected from a grassland in the Netherlands, in spring 2013 (‘Clue’ site, Mosselse Veld, 52 040 N, 5 450 E). The soil is sandy-loam, with particle size distribution: <2 mm, 3.4%; 2e63 mm, 17.3%; >63 mm, 79.7%, pH H2O 6.4 and %OM 4.5 (Van der Putten et al., 2000; Bezemer et al., 2010). Prior to filling the pots part of the soil was sterilised by gamma irradiation (25 kGray). The unsterilized soil was stored at 4  C for two weeks until the soil sterilisation was completed. With the sterilised and living soil a soil mixture was prepared consisting of 85% sterilised soil and 15% living soil from which the plants could develop their own soil community. We sterilised part of the soil in order to avoid weed pressure and insect root herbivores but inoculated with living soil to provide a natural pool of soil microbes. Plants were surface sterilised by dipping them in diluted household chloride bleach (10% bleach solution for 30s), and afterwards rinsed several times with tap water. The seeds were subsequently germinated on autoclaved sand and two-week old seedlings were transplanted individually to 2 L pots filled with the soil mixture and grown in the greenhouse for 12 weeks (with day:night regime of 16:8 h light:dark, 21:16  C). At harvest, the shoots were separated from the roots. The roots were washed carefully and dried at 40  C for one week. The roots were weighed and a subsample of fine roots (diameter < 2 mm) was

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collected to be used in the litter bags. The roots of two independently grown plant individuals of each species were homogenized by cutting them into 5 mm long pieces; thus creating an independent root litter pool to be used in the experiment. This procedure was repeated four times for each species, creating four independent root litter pools per species, each containing roots of two plant individuals. Each root litter pool was used to fill four litterbags, each containing 0.5 g of dried roots; two of these litter bags were used for the treatment without living plant and two for the treatment of decomposition with living plant. Per litter species there were 16 litterbags in total, eight to decompose without and eight to decompose with the presence of a white clover plant. The litter bags were made of polyester fabric (mesh size < 0.05 mm, litter bag size 70  70 mm stitched at 5 mm from the edge and closed after filling by using three staples. The mass of the empty and full litter bag and of the root litter in the litter bag was recorded.

2.3. Soil and plant nutrient analysis We measured mineral plant available N and P in the soil using the standard CaCl2 extraction method (Houba et al., 2000). We quantified the concentrations of N, P and C in the root litter prior to incubation for each root species on a subsample of each root litter pool comprising two individual plants, resulting in four independent litter samples for chemical analysis per species. The subsamples of dried roots were ground, then mineralised using H2SO4/ H2O2/Se wet digestion (Novozamsky et al., 1983), and subsequently analysed for total N and P content using Segmented Flow Analyses (SFA). Another subsample of ground roots was used to quantify the C and N concentration in the litter using a CN elemental analyser (LECO, Germany); due to limited litter availability for this CN analyses there were three instead of four replicates for the species L. perenne, T. pratense and V. cracca and two for T. repens. 2.4. Data analysis

2.2. Litter decomposition experiment The experiment combined two treatments: 1) the presence of a growing plant of T. repens (two levels: absence, presence) and 2) root litter species (seven levels: root litter of seven different plant species) and was conducted in the greenhouse during January to May 2014. The number of replicates per treatment of litter with and without plant presence per litter species was eight. Prior to germination, the seeds of T. repens were surface sterilised in the same way as described above and stored overnight at 4  C. The experimental soil was a mixture of 10% living soil and 90% sterilised soil (same soil type and origin as described above). The soil nutrient levels in the soil mixture were: 85.8 ± 5.0 PePO4, 270.6 ± 15.0 NeNH4, 91.4 ± 15.0 N-(NO3 þ NO2) mg per kg soil dry weight, determined using standard methods (see below). The seeds were geminated in a greenhouse in plastic cups (200 ml) filled with experimental soil. Eight-week-old plants were transferred into pots (11  22  12 cm) filled with 1740 g of the same soil with moisture level at 60% of the water holding capacity (WHC). The soil surface was covered with autoclaved sand to prevent growth of mosses and algae. The pots were kept in the greenhouse at 16/21  C (night/day) with additional light when needed to achieve a 16 h day length and watered daily to 60% WHC by adjusting the pots by adding water to their target initial weight. The pots were placed in the greenhouse in a complete randomised block design with eight blocks. Each root litter pool of each species was represented in two of the total of eight blocks, with a replicate per treatment per block. The litterbags were inserted into all pots ten days after planting the clover plants. The litterbags were placed vertically at 2 cm from the pot border and the same positioning was used in the pots without plants. Three weeks after planting leaves and protruding stems of all clover plants were clipped to 2 cm height to ensure active plant (re)growth during the experiment and to prevent the plants from growing in neighbouring pots. After eight weeks of decomposition the plants were harvested, shoots and roots were separated and roots were rinsed from soil and root nodulation was confirmed in all harvested plants. Shoots and roots were dried at 40  C for 48 h and weighed. The litter bags were recovered from the soil during the harvest, rinsed and dried under the same conditions as the plant material. The remaining litter was collected from within the dried litter bag and weighed. A decomposition period of eight weeks is relatively short. However, the temperature in the greenhouse was favourable for decomposition and litter mass loss during this period was in the range of 30e80%, which enabled testing of treatment effects.

The decomposition rate of the roots was calculated for each litter bag as the proportion of litter mass that was lost during the eight weeks of the experiment:

Litter loss ¼ ð1  ðLitter massend =Litter massstart ÞÞ Priming effect was calculated for each litter species as the difference between the mean root decomposition rate in the presence versus in absence of a living plant in the soil:

  Priming effect ¼ Litter losswith plant  Litter losswithout plant For each litter species Litter loss with plant and Litter loss without plant were averaged for each independent root litter pool (n ¼ 4) before calculating the priming. To test for species-specific root decomposition patterns in relation to plant presence, a linear mixed model was fitted with the proportion of litter decomposed (Litter loss) as a response variable and litter species, presence/absence of the living plant and their interaction as predictor variables. Root litter pool (i.e. litter pool 1 to 4) and block nested within root litter pool were included as random factors in the model. Decomposition rate data were arcsine and square root transformed before the analysis. To test for the effects of litter N, P and C concentrations and C:N ratio on litter decomposition rate, linear mixed models were designed with priming effect or litter decomposability (i.e. Litter loss in the absence of living plants) as a response variable and concentrations of N, P, C or C:N as a predictor variable and litter species as a random factor. Since N, P, C and C:N concentration were measured for each species and each root litter pool (n ¼ 4 per litter species), the priming effect and decomposability were also averaged across each litter species and each root litter pool before the analysis. F-statistics and residual degrees of freedom in mixed models were calculated using Kenward-Roger approximation. Data analyses were performed using R (version 3.1.0) packages lme4 (Bates et al., 2014) and multcomp (Hothorn et al., 2008). 3. Results 3.1. Interactive effects of plant presence and root litter species on decomposition rate The effect of plant presence on root litter mass loss depended significantly on the species of the decomposing root litter (interaction plant presence x root litter species: F6,91 ¼ 14.7, P < 0.001; Table 1). Overall the plant species differed in decomposition rate

S. Saar et al. / Soil Biology & Biochemistry 94 (2016) 88e93 Table 1 Root litter mass loss (%) after eight weeks of decomposition in relation to litter species and the absence or presence of a growing Trifolium repens plant (means ± SE, n ¼ 8). Asterisks indicate significant differences between treatments within a litter species: Ns not significant, *P < 0.05, ***P < 0.001. The significance test was performed on arcsin sqrt transformed % mass loss data and was adjusted for multiple comparisons using a single-step method. Root litter species

Without T. repens

Cichorium intybus Vicia cracca Trifolium repens Trifolium pratense Arrhenatherum elatius Lolium perenne Festuca rubra

79.7 70.1 58.9 57.8 56.4 41.0 36.5

± ± ± ± ± ± ±

0.5 1.4 1.3 0.7 1.3 2.1 1.2

With T. repens 75.9 71.6 61.8 58.8 43.4 29.9 31.6

± ± ± ± ± ± ±

0.8 1.4 1.2 1.2 0.9 1.4 1.0

P * Ns Ns Ns *** *** *

with root litter of C. intybus decomposing the fastest (more than 75% mass loss in eight weeks) and root litter of F. rubra decomposing the slowest (less than 40% mass loss, Table 1). The ranking of the species from the fastest to the slowest decomposing roots was the same irrespective of the presence of T. repens. However, the presence of a growing clover plant significantly reduced the decomposition rate of the roots of all non-legume species, especially in the relatively slow decomposing grasses A. elatius and L. perenne (P < 0.001) and F. rubra (P ¼ 0.014) but also in the fast decomposing forb C. intybus (P ¼ 0.039). In contrast, the root decomposition rate of the three legume species was not significantly affected by the presence of a growing clover plant (P > 0.05; Table 1). Root litter species had no significant effect on the growth of white clover plants: both aboveground biomass (F6,42 ¼ 0.28, P ¼ 0.943) and belowground biomass (F6,42 ¼ 0.72, P ¼ 0.639) remained unaffected. 3.2. The effect of root litter quality on priming and decomposability The priming effect (the difference in litter mass loss in the presence versus in the absence of a living plant) showed a

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significant positive relation with the percentage of N (F1,6 ¼ 13.8, P ¼ 0.011; Fig. 1a) and a positive trend with the percentage of P (F1,14 ¼ 3.1, P ¼ 0.099; Fig. 1b) in the initial root litter, indicating that the priming effect was less negative the more N and P rich the initial root litter was. Moreover we found a significant negative relation between the priming effect and litter C:N ratio (F1,6 ¼ 8.3, P ¼ 0.027; Fig. 1d) and no relation with initial root C concentration in the decomposing roots (F1,16 ¼ 0.12, P ¼ 0.734; Fig. 1c). Decomposition of root litter of non-legume species, which were characterised by low N and P concentrations and by a high C:N ratio, was slowed down in the presence of the growing clover plant, while the root litter decomposition rate of the legume species was not altered by the presence of white clover. When the data was split to legume and non-legume species, no significant relationship between priming effect and root litter properties (N%, P%, C% and C:N) was found within legume and non-legume species groups (P > 0.05). In contrast to the priming effect, litter decomposability (i.e. % root litter mass loss in the absence of a growing clover) did not depend on the initial concentration of N (F1,15 ¼ 0.02, P ¼ 0.891; Fig. 1e), P (F1,25 ¼ 0.91, P ¼ 0.349; Fig. 1f), C (F1,15 ¼ 0.84, P ¼ 0.373; Fig. 1g), or C:N (F1,18 ¼ 0.01, P ¼ 0.922; Fig. 1h) of the decomposing root species. 4. Discussion The main goal of this study was to test whether the rate of root litter decomposition is affected by the presence of a living plant and whether the effect depends on the initial chemical composition of the root litter. We hypothesised that the presence of a living plant of T. repens would increase the decomposition rate of the roots across species, and that the magnitude of such a priming effect would depend on the nutrient concentrations in the decomposing roots. We did not find the expected acceleration of root decomposition with the presence of a living clover plant; instead, we found suppression or no change in decomposition rate

Fig. 1. Relation between root litter properties and priming effect of root litter mass loss (aed) and root litter mass loss (eeh). Priming effect in relation to root litter N concentration (F1,6 ¼ 8.579, P ¼ 0.0254) (a), P concentration (F1,14 ¼ 3.124, P ¼ 0.099) (b), C concentration (F1,16 ¼ 0.120, P ¼ 0.734) (c) and C:N ratio (F1,6 ¼ 8.252, P ¼ 0.027) (d). Priming effect was calculated as the difference between litter mass loss rates in the presence versus absence of a living plant. Negative values signify negative priming i.e. slower litter decomposition in presence of a living plant. Root mass loss (% mass loss after 8 weeks in the absence of a living plant) in relation to root litter N concentration (e), P concentration (f), C concentration (g) and C:N ratio (h). Open circles denote non-legumes and filled circles denote legumes. Ci ¼ Cichorium intybus, Vc ¼ Vicia cracca, Tr ¼ Trifolium repens, Tp ¼ Trifolium pratense, Al ¼ Arrhenatherum elatius, Lp ¼ Lolium perenne, Fr ¼ Festuca rubra; n ¼ 4 per species except for C:N and C where n ¼ 3 for Lp, Vc and Tp and n ¼ 2 for Tr. *P < 0.05; $ P < 0.1; NS P > 0.1.

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depending on the root litter species. Root decomposition rate of non-legume root litters was suppressed by the presence of a growing clover plant (i.e. negative priming effect) and these litters were characterised by low initial concentrations of N and P. In contrast, the decomposition of legume root litters, of Trifolium as well as of Vicia species, was unaffected by the growing clover presence; these legume root litters were consistently richer in N and P and had a low C:N ratio. The effect of T. repens on the decomposition rate of roots of Trifolium pratensis and V. cracca appeared to be similar, while a trend of accelerated root decomposition of T. repens roots could be noted. This trend may be due to some home-field advantage, by which root litter of T. repens decomposes faster in the presence of a plant of its own species (Ayres et al., 2009; Freschet et al., 2012a). However, the potential presence of a home-field advantage cannot explain the suppression of root decomposition rates in the nonlegume species as those species should not be affected in their decomposition by the presence of a non-related species. Future studies will benefit from combining tests of priming effects induced by living plant species with home-field advantage tests by manipulating the species identity of both the living plant and root litter as factors in the experimental design. To date, most studies have found enhanced rates of decomposition, i.e. positive priming effects, due to the presence of living plants, as reviewed by Cheng et al. (2003) and shown by later studies (Phillips and Fahey, 2006; Dijkstra and Cheng, 2007; Zhu et al., 2011; Bengtson et al., 2012; Phillips et al., 2012). However, some studies have found negative priming effects due to the presence of some plant species (Van der Krift et al., 2002; Coq et al., 2011), in line with our results. These latter studies explained negative priming effects by indirect effects of growing plants on soil biota, namely through soil moisture fluctuations which were larger with than without a growing plant (Sparling et al., 1982; Smith et al., 2014). In our study, we watered the pots daily and adjusted the moisture of both the planted and unplanted pots very regularly to the standardised level of 60% WHC. Moreover, we found interactive effects between the presence of a growing clover plant and the root litter species with no priming effect on root litter of legume species, while aboveground biomass of the living plant, and hence likely also water use, was not altered by the identity of the root litter species. Therefore, we think that the role of different soil microclimate with respect to soil moisture cannot explain our results. Negative priming effects can also appear when mineral N-concentration in soil is low and plants are competing with soil microbes for N (Van der Krift et al., 2002; Bardgett et al., 2003). This is a plausible mechanism in our case because the soil we used was relatively nitrogen poor and the plants showed signs of N and K deficiency as evidenced by mild chlorosis and small brown spots (McNaught, 1958). The N:P ratio of the roots of T. repens grown on this soil was 4.6e7.1 indicating that, despite being a legume and able the fix atmospheric nitrogen, T. repens plants were limited by N rather than by P availability (Almeida et al., 2000). However, root litter contained a small proportion of the total nutrient pool in the soil as only 0.5 g of dry root litter was added to each pot in our experiment, which may explain why the plant biomass did not differ between root litter treatments. Given that the plants were similar in size across litter treatments and that the soil provided most of the nutrients, it is unlikely that the intensity of competition between plants and soil microbes was strongly affected by differences in N content between root litter species and caused a negative priming effect, unless if only very locally around the N-poor litter. An alternative mechanism for negative priming effects is formulated in the Preferential Substrate Utilization (PSU)

hypothesis, which assumes that when soil organic matter requires too much energy to be degraded, microorganisms switch to use fresh, easily decomposable organic matter such as root exudates as a C source (Cheng and Kuzyakov, 2005). Rhizodeposition by legumes is also known to increase N availability in the soil, even when N is limiting the growth of legumes themselves (Fustec et al., 2011). According to the microbial N mining theory (Craine et al., 2007), N added to the soil by the roots of T. repens could have shifted microbial activity away from recalcitrant litter, which is being decomposed for its N content when N is scarcely available. It has been shown that N-addition to soil especially reduced the decomposition of litters consisting of more recalcitrant C compounds (Craine et al., 2007). Our results are in line with this idea as grass roots generally contain more recalcitrant carbon forms than legume roots (Birouste et al., 2012) and we found the strongest negative priming effects in the grass species and the weakest priming effects in legumes. However, detailed measurements of root exudation profiles and carbon and nitrogen utilisation by soil biota are necessary to test this possible mechanism of negative priming. We found that the strength of the priming effect depended on root litter N and P concentrations, showing the most negative priming in low-quality litter, i.e. poor in N and P. The litter nutrient concentration in N and P can explain differences in the priming effect between legume versus non-legume litter; however, it cannot explain the variation in priming effect among non-legumes. Also, the decomposition rate of the root litters in the absence of a living plant could not be explained by root litter N, P and C concentration or C:N ratio. Therefore, our results cannot be entirely explained by nutrient concentrations and other litter characteristics must be involved. Although total C quantity in the dead roots did not explain differences in decomposition rates, there may be differences in C quality between our root litter species that may affect the decomposability and the level of priming effect by living plants. Several studies pointed to the role of lignin concentration and lignin:N ratio as important litter traits for explaining litter decomposability (Osono and Takeda, 2004; Vivanco and Austin, 2006; Bontti et al., 2009). However, these studies are based on longer-term incubation, often examining woody plant species. All our species appeared to be fairly easy to decompose given that after eight weeks the litter mass loss ranged from 30% to 80%. Moreover, our study focused on the first phase of litter decomposition in which easily decomposable C compounds, rather than structural compounds like lignin, are thought to be important in determining the speed of decomposition (Aerts, 1997; Freschet et al., 2012b). Thus, further research is necessary to disentangle the complex relationship between the strength of the priming effect, differences in concentrations of labile and non-labile C in the litter and N availability (Craine et al., 2007). In conclusion, we found that the presence of living legume roots can decrease the rate of root litter decomposition and that this reduction can in part be explained by the root litter N and P concentrations, with greater suppression of decomposition for root litter poor in N and P. Differences in priming effects between non-legume species warrant yet other litter properties to be identified as a driving factor. The negative priming effect found for non-leguminous litter may have been caused by preferential substrate utilisation, whereby soil biota switch to consuming easily decomposable root exudates instead of decomposing poorquality root litter. Moreover, N deposition in the clover rhizosphere may have inhibited microbial N mining from the litter with low N concentration. Further studies involving simultaneous manipulation of root exudate and litter quality are necessary next steps in testing these possible mechanisms behind negative priming effects.

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Acknowledgements We thank Henk Martens for help in root washing, Elly Nijenhuis for sewing the litter bags, and Jaap Nelemans, Gerlinde Vink and CLBB colleagues for laboratory assistance. This research was supported by NWO-ALW Vidi (grant 864.11.003) granted to GB De Deyn, Estonian national scholarship program Kristjan Jaak granted to S. Saar (Archimedes Foundation in collaboration with the Ministry of Education and Research in Estonia), targeted financing (IUT 20-31) and Estonian Science Foundation (grant 9332) granted to M. Semchenko. References Aerts, R., 1997. Climate, leaf litter chemistry and leaf litter decomposition in terrestrial ecosystems: a triangular relationship. Oikos 79, 439e449. €sberger, J., Lüscher, A., 2000. Evidence Almeida, J.P.F., Hartwig, U.A., Frehner, M., No that P deficiency induces N feedback regulation of symbiotic N2 fixation in white clover (Trifolium repens L.). Journal of Experimental Botany 51, 1289e1297. 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