Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids

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Pedobiologia - Journal of Soil Ecology journal homepage: www.elsevier.de/pedobi

Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids

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Andrea Grabmaier a , Florian Heigl a , Nico Eisenhauer b,c , Marcel van der Heijden d , Johann G. Zaller a,∗ a

Institute of Zoology, University of Natural Resources and Life Sciences Vienna, Gregor Mendel Straße 33, A-1180 Vienna, Austria German Centre for Integrative Biodiversity Research, University of Leipzig, Deutscher Platz 5e, 04103 Leipzig, Germany c Institute for Biology, University of Leipzig, Johannisallee 21, 04103 Leipzig, Germany d Agroscope, Reckenholzstrasse 191, CH-8049 Zurich, Switzerland b

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a r t i c l e

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a b s t r a c t

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Article history: Received 27 May 2014 Received in revised form 20 October 2014 Accepted 20 October 2014

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Keywords: Aboveground–belowground interactions Aphids Arbuscular mycorrhizal fungi Earthworms Multitrophic interactions Stable isotopes

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Introduction

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Functional relationships between belowground detritivores and/or symbionts and aboveground primary producers and their herbivores are not well studied. In a factorial greenhouse experiment we studied interactions between earthworms (addition/no addition of Lumbricus terrestris; Clitellata: Lumbricidae) and arbuscular-mycorrhizal fungi (AMF; with/without inoculation of Glomus mosseae; Glomerales: Glomeraceae) on the leguminous herb Trifolium repens (Fabales: Fabaceae) and associated plant aphids (Aphis gossypii, A. craccivora; Hemiptera: Aphidoidea). In order to be able to trace organismic interactions, earthworms were dual-labelled with stable isotopes (15 N-ammonium nitrate and 13 C-glucose). We specifically wanted to investigate whether (i) isotopic signals can be traced from the labelled earthworms via surface castings, plant roots and leaves to plant aphids and (ii) these compartments differ in their incorporation of stable isotopes. Our results show that the tested organismic compartments differed significantly in their 15 N isotope enrichments measured seven days after the introduction of earthworms. 15 N isotope incorporation was highest in casts followed by earthworm tissue, roots and leaves, with lowest 15 N signature in aphids. The 13 C signal in roots, leaves and aphids was similar across all treatments and is for this reason not recommendable for tracing short-term interactions over multitrophic levels. AMF symbiosis affected stable isotope incorporation differently in different subsystems: the 15 N isotope signature was higher below ground (in roots) but lower above ground (leaves and aphids) in AMF-inoculated mesocosms compared to AMF-free mesocosms (significant subsystem × AMF interaction). Aphid infestation was unaffected by AMF and/or earthworms. Generally, these results demonstrate that plants utilize nutrients excreted by earthworms and incorporate these nutrients into their roots, leaf tissue and phloem sap from where aphids suck. Hence, these results confirm that earthworms and plant aphids are functionally interlinked. Further, 15 N-labelling earthworms may represent a promising tool to investigate nutrient uptake by plants and consequences for aboveground multitrophic interactions. © 2014 Published by Elsevier GmbH.

In the last decade it has increasingly been recognized that a combined aboveground–belowground approach is necessary to understand the functioning of terrestrial ecosystems (Wardle et al., 2004; van der Putten et al., 2009; Bardgett and Wardle, 2010; Eisenhauer, 2012). Plants thereby play an essential role as they

∗ Corresponding author. Tel.: +43 1 47654 3205; fax: +43 1 47654 3203. E-mail address: [email protected] (J.G. Zaller).

interlink above- and belowground subsystems. Factors above the soil surface can directly or indirectly influence the plant itself, but can also affect soil processes and soil organisms that can feed back to plants (Bardgett and Wardle, 2003; Porazinska et al., 2003; Schröter et al., 2004; Wardle et al., 2004; van der Putten et al., 2009). Several studies investigating the functional diversity and multitrophic interactions in terrestrial ecosystems have shown that aboveground–belowground interactions can have consequences at the ecosystem level (Scheu, 2001; Wardle et al., 2004; Megías and Müller, 2010; Eisenhauer and Schädler, 2011; Zaller et al., 2011b; Eisenhauer, 2012; Arnone et al., 2013). However, so far only a few

http://dx.doi.org/10.1016/j.pedobi.2014.10.002 0031-4056/© 2014 Published by Elsevier GmbH.

Please cite this article in press as: Grabmaier, A., et al., Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids. Pedobiologia - J. Soil Ecol. (2014), http://dx.doi.org/10.1016/j.pedobi.2014.10.002

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studies have focussed on the effects of belowground detritivores and symbionts on aboveground herbivory (e.g. Poveda et al., 2003; Megías and Müller, 2010; Wurst, 2010; Wurst and Rillig, 2011; Trouvé, 2013). Earthworms make up the majority of the soil faunal biomass in temperate grasslands (Lee, 1985; Curry, 1994) and are considered as ecosystem engineers (e.g. Jones et al., 1994). They alter the quality of resource inputs either directly, through the return of up to 45 ton ha−1 a−1 of nutrient rich casts (Bohlen et al., 1997), or indirectly by altering soil processes through bioturbation, acceleration of decomposition of organic materials and an increase in microbial activity and nutrient mineralization, which results in increased plant nutrient uptake and plant growth (Brussaard, 1999; Bonkowski et al., 2001). Despite the plethora of information on the effects of earthworms on soil structure, nutrient availability and plant growth (Bohlen et al., 1997; Scheu, 2003), little is known about their effects on aboveground herbivores (Wurst, 2010). A few studies report earthworm-stimulated herbivory (Scheu et al., 1999; Wurst and Jones, 2003; Poveda et al., 2005), but it appears that no effects or negative effects of earthworms on aboveground herbivores are more often found (Bonkowski et al., 2001; Wurst et al., 2003; Wurst et al., 2004; Trouvé, 2013; Zaller et al., 2013a). Arbuscular mycorrhizal fungi (AMF) form a widespread mutualism between the plant roots of over 80% of all families of land plants (Smith and Read, 2008). In most environmental conditions, these fungi are beneficial to their host plants, by providing access to limiting soil nutrients, increasing photosynthetic rates and resistance to drought, insect herbivores and fungal pathogens (reviewed in van der, 2002; Smith and Read, 2008). Studies on the effects of AMF on herbivores show either positive or no effects of AMF on phloem feeding insects (Koricheva et al., 2009; Reidinger et al., 2012). In most terrestrial ecosystems of the temperate region, earthworms and arbuscular mycorrhizal fungi are commonly co-occurring and interacting with each other, however our understanding of these functional interactions is still very rudimentary. Generally, earthworms are thought to affect AMF populations by (i) ingesting fungal species (Dash et al., 1979; Cooke, 1983; Edwards and Fletcher, 1988; Morgan, 1988; Kristufek et al., 1992; Bonkowski et al., 2000) thus affecting the germination of ingested spores (Parle, 1963; Hoffmann and Purdy, 1964; Keogh and Christensen, 1976; Striganova, 1988) and (ii) dispersal of AMF spores (Coûteaux, 1994; Pattinson et al., 1997; Wurst et al., 2004). Even less is known about the effects of earthworms and AMF on aboveground herbivores, especially on sap-sucking herbivores such as aphids. In both natural and agricultural ecosystems aphids are known to affect plant growth, biomass production, phenology and chemistry (Dixon, 1998). The presence of both earthworms and AMF was shown to accelerate the development of an aphid species (Myzus persicae), while aphids were delayed when only AMF or earthworms were present (Wurst et al., 2004). In another study aphid abundance on plants decreased in treatments with earthworms, but this was independent of the presence of AMF (Wurst and Forstreuter, 2010). Despite these important contributions, the mode of interaction between earthworms, AMF, plants and sap-feeding insects remains uncertain and mainly conceptual as traditional ecological methods provided only limited information on nutrient fluxes among organisms involved in these trophic relationships. To better understand potential multitrophic interactions between earthworms and other organisms in terrestrial ecosystems we for the first time used 15 N–13 C dual stable isotope labelling of earthworms (Schmidt et al., 2004; Heiner et al., 2011) in experimental ecosystems comprising AMF, plants and aphids. We used 15 N ammonium-nitrate and 13 C glucose as both elements can be taken up by plant roots either directly or when incorporated in amino acids or other organic sources (Marschner, 1995). For the

present study, we hypothesized that due to functional interactions the isotopic label can be detected in different parts of this model ecosystem and that AMF would increase stabile isotope uptake into plants and aphids. In particular we investigated whether (i) isotopic signals would be passed on from labelled earthworms to surface castings, plants and aphids, and (ii) these compartments differ in their incorporation of stable isotopes. These objectives were tested in a factorial experiment where we manipulated the factors earthworms (Lumbricus terrestris) and AMF (Glomus mosseae) and studied their single or interactive effects on the legume Trifolium repens and associated aphids (Aphis spp.). The chosen species are commonly co-occurring and interacting in temperate grassland ecosystems throughout Europe.

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Materials and methods

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Experimental setup and treatments

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The experiment was carried out in the research greenhouse of the University of Natural Resources and Life Sciences, Vienna (BOKU) between March and November 2011. Using a full-factorial design we manipulated earthworms (two levels – addition of L. terrestris, +EW, without earthworms, −EW) and AMF inoculation (two levels – inoculation with G. mosseae, +AMF, no AMF inoculation, −AMF) to investigate interactions between earthworms, AMF, plants and aphids. We set up six replicates of each treatment totalling 24 experimental units. Experimental units consisted of 20 L plastic planting pots (diameter: 31 cm, height: 30 cm; further called mesocosms) filled with 18 L sterilized field soil (Haplic Chernozem, silty loam; steamed at 100 ◦ C for 15 h) and quartz sand (grain size 1.4–2.2 mm) in a ratio of 40:60 (v/v) (bulk density 1.4 g cm−3 , pH = 7.6, Corg = 22.0 g kg−1 , Ntot = 0.92 g kg−1 , P-CAL = 64.5 mg kg−1 , K-CAL = 113.6 mg kg−1 ). We successfully used this substrate mixture in other experiments involving the same earthworm, plant and AMF taxa (e.g. Putz et al., 2011; Zaller et al., 2011c; Zaller et al., 2013b). All mesocosms were lined at the bottom with two layers of garden fleece material to prevent earthworms from escaping while still allowing water drainage. The uppermost 10 cm soil layer (approximately 6 L) was inoculated with AM fungal propagules [Glomus mosseae (T.H. Nicolson & Gerdemann) Gerdemann & Trappe (La Banque Européenne des Glomales – BEG 198) (Glomeraceae)] by mixing 25 g L−1 of commercial AMF inoculum (Symbion, Landskroun, Czech Republic) to the substrate (treatment +AMF). AMF-free control treatments (−AMF) received the same amount of sterilized, inactive AMF inoculum (steamed at 110 ◦ C for 2 h). In addition, each mesocosm received 100 ml microbial wash from active AMF inoculum and 300 ml microbial wash of field soil. This microbial wash corrects for possible differences in microbial communities between the different treatments (Koide and Li, 1989). It was prepared with a total of 0.5 kg AMF inoculum and 3.5 kg field soil that was wet-sieved through a cascade of sieves, where the finest sieve had a mesh size of 25 ␮m to receive a filtered non-mycorrhizal microbial inoculum. Furthermore, Trifolium repens L. (Fabaceae) seeds from an agricultural seed supplier (Lagerhaus, Groß-Enzersdorf, Austria) were germinated in commercial steam-sterilized (105 ◦ C for 20 h) potting soil on a greenhouse bench. Eight days after germination, 18 seedlings (approx. 2 cm high) were transplanted into each mesocosm in a consistent hexagonal pattern in equal distance of each plant individual of 5 cm (equals 240 plants m−2 ). The greenhouse containing the mesocosms was not temperature-controlled, artificial light was only provided during the last 38 days (14 h light day−1 in October and November). Mean air temperature during the course of the experiment was 21.7 ◦ C at 51.6% mean relative

Please cite this article in press as: Grabmaier, A., et al., Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids. Pedobiologia - J. Soil Ecol. (2014), http://dx.doi.org/10.1016/j.pedobi.2014.10.002

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humidity. The mesocosms were irrigated daily by adding 500 ml of demineralised water mesocosm−1 and were randomly rearranged every two weeks to avoid a bias of potential environmental gradients within the greenhouse. No fertilizers were applied during the course of the experiment.

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Isotopic labelling of earthworms

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In order to be able to trace the interactions between earthworms, AMF, plants and aphids, earthworms were labelled with 15 N and 13 C stable isotopes prior to introduction into the respective mesocosms. Adult L. terrestris (Linnaeus 1758) were obtained from a fishing bait shop (Anglertreff E. Lux, Vienna, Austria). Prior to the labelling, all earthworms were cultivated in sterile soil and fed with ground oat flakes for four days. Following (Heiner et al., 2011) we added 100 mg of 13 C6 H12 O6 (99 at.% 13 C6 -glucose; Campro Scientific, Berlin, Germany) and 100 mg 15 NH4 NO3 (98 at.% 15 N-ammonium nitrate; Campro Scientific, Berlin, Germany) to 200 g of dry field soil and stored it in a climate chamber at 15 ◦ C for eleven days. After this incubation time earthworms from sterile soil were carefully rinsed with water, transferred into the labelled soil mixture and kept there for seven days to allow proper incorporation of 15 N and 13 C into their tissue and gut. We placed two adult earthworm individuals per 100 g of soil-isotope mixture; earthworms were not fed during this labelling period. Two months prior to the end of the experiment, four labelled adult specimens of L. terrestris with a mean initial biomass of 3.89 ± 0.14 g per individual (mean ± SE) were added to each mesocosm of the +EW treatment, resembling a density of 204.8 g m−2 , which is slightly higher than the ambient density in permanent grasslands (Decaens et al., 1997). The upper rim of all mesocosms was extended with a transparent plastic barrier (10 cm height) to prevent earthworms from escaping.

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Aphids

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Aphids (Aphididae) coming into the greenhouse through the permanently open side windows colonized the mesocosms starting about eleven weeks prior to the insertion of the labelled earthworms. Aphids comprised a mixed abundance of 50% Aphis gossypii (Glover 1877) and 50% Aphis craccivora (Koch 1854; E. Koschier, personal communication). Aphid abundance was estimated once a week over a period of 10 weeks. The investigation of freely moving aphids has been proven successful in exploring aphid selectivity for host plants and reproduction in response to manipulation of soil organisms (Eisenhauer and Scheu, 2008). Sampling and analytical measurements Aboveground plant growth was assessed on days 71, 115 and 189 after germination by measuring plant height from the soil surface to the uppermost end of the leaves. Plant biomass production was assessed by cutting the vegetation 1 cm above the soil surface on the dates mentioned above. For the statistical analyses the cumulated height growth and cumulated biomass production (sums of the three measurements and harvests) were used. Plant material was dried at 50 ◦ C for two days and weighed to determine dry mass production. Samples for 13 C and 15 N analyses were taken one week after the introduction of the labelled earthworms by randomly collecting two surface casts per mesocosm, clipping five Trifolium leaves per mesocosm and collecting approximately 80 aphids per mesocosm; three root samples were taken using a corer (diameter: 3 cm, depth: 20 cm) inserted near Trifolium plants three weeks after earthworm introduction. Plant leaves and earthworm casts were dried at 60 ◦ C for 24 h, aphids and root samples were freeze-dried

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for 48 h. Subsequently all samples were ground using a ball mill. Samples for 13 C and 15 N analyses in earthworms were taken 58 days after their introduction by collecting one earthworm per mesocosm. This was done at night when earthworms were searching for food on the soil surface. Collected earthworms carefully rinsed in distilled water and weighed. For isotopic analysis of the earthworm tissue only the anterior 15 segments of the earthworms without gut content were used, this tissue part was freeze-dried and homogenized manually using a mortar and pestle. For isotopic analysis 1.0 mg of aphids, 1.8 mg of leaf material, 1.8 mg of root material, 5.0 mg of earthworm casts, and 1.0 mg of earthworm tissue samples were weighed into tin capsules and analysed for N and C by continuous flow isotope ratio mass spectrometry (CF-IRMS) at the SILVER lab of the Department of Terrestrial Ecosystem Ecology at the University of Vienna, Austria. Ratios (R) of 15 N: 14 N and 13 C: 12 C were expressed relative to the international standards (atmospheric air for 15 N and Vienna Pee Dee Belemnite for 13 C) in parts per thousand (␦‰) using the formula Rsample × Rstandard −1 × 1000. The analytical precision was 0.15‰ for N and 0.1‰ for C isotopes, respectively. Natural abundance values of 15 N and 13 C of earthworm tissue and casts were obtained from earthworms that were cultivated in the same way as described above, except no isotopic labels were added to the substrate. To quantify AMF root colonization rates fresh root subsamples were cleared with boiling KOH and vinegar for 4 min and stained for 1 min with Shaeffer® black ink (Vierheilig et al. 1998). Stained roots were analysed under a dissecting microscope (100× magnification) and the percentage of colonized roots with AMF was calculated using the gridline intersection method by counting 200 intersections per sample (Giovanetti and Mosse, 1980). Statistical analysis Normal distribution (Shapiro–Wilk test) and homogeneity of variances (Levene’s test) were tested and the data were transformed if necessary to match the prerequisites for analysis of variance (ANOVA). Multiple samples per mesocosm were averaged to create one value per mesocosm. One-way ANOVAs were performed to test for effects of earthworms on root mycorrhization and for effects of AMF on earthworm numbers and biomass, on cumulated plant height growth, on cumulated aboveground biomass production and on aphid abundance. Two way ANOVAs with AMF and earthworms as factors were performed to test for treatment effects on 13 C and 15 N in earthworm tissue, earthworm casts, roots, leaves and aphids. Two way ANOVAs were also performed to test for effects of earthworms and AMF on 13 C and 15 N signatures of the aboveground subsystem (comprising leaves and aphids) vs. the belowground subsystem comprising roots. Significance of differences between means of two or more groups was analysed using Tukey’s HSD post hoc tests. Linear regressions were performed to explore relationships between isotopic enrichments of the different compartments. All statistical analyses were conducted using the software IBM SPPS Statistics 20 (SPSS Inc. Headquarters, Chicago, Illinois, USA). Values given throughout the text are means ± SD.

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Results

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Mycorrhiza and earthworms

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Across the AMF-inoculated mesocosms 32.3 ± 1.6% of the roots of T. repens were colonized by AMF; 3.6 ± 2.3% of the roots of the AMF-free treatments were colonized by AMF. Percent root length colonized by AMF was significantly decreased in the presence of earthworms (−EW = 34.0 ± 1.6%, +EW = 29.0 ± 2.0%; F1,22 = 4.74, P = 0.042). At the end of the experiment, the biomass of the

Please cite this article in press as: Grabmaier, A., et al., Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids. Pedobiologia - J. Soil Ecol. (2014), http://dx.doi.org/10.1016/j.pedobi.2014.10.002

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Fig. 1. Isotopic enrichment of ␦15 N in (a) earthworm surface casts, (b) earthworm tissue, (c) aphids, (d) leaves and (e) roots of Trifolium repens in mesocosms without (−AMF) and with (+AMF) inoculation with arbuscular mycorrhizal fungi. Note different scales on y-axes. Means ± SD, n = 5–6.

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recaptured earthworms was 3.53 ± 0.23 g earthworm−1 ; a decrease of 9.2% compared with the initial weights. Neither earthworm numbers at the end of the experiment (F1,8 = 0.39, P = 0.55) nor recaptured earthworm biomass were affected by AMF (F1,12 = 0.01, P = 0.91).

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Plant performance and aphid infestation

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Both cumulated height growth (+21%) and cumulated aboveground biomass (+19%) production of T. repens was significantly increased in the presence of AMF (height: +AMF = 15.4 ± 1.0 cm: F1,22 = 50.68, −AMF = 12.9 ± 0.9 cm; P < 0.001; biomass: −AMF = 9.3 ± 1.4 g; +AMF = 11.0 ± 2.3 g: F1,22 = 5.01, P = 0.036). The abundance of aphids was not significantly affected by AMF treatments (F1,22 = 2.18, P = 0.15; total number of aphids mesocosm−1 : −AMF = 663.3 ± 75.0, +AMF = 781.7 ± 59.5). No statements about earthworms effects on plant growth, plant production and aphid abundance can be made, because earthworms were inserted into the mesocosms after the third biomass harvest and aphid abundance estimations were not continued during this time.

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Isotopic enrichments and nutrient concentrations

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Mean ␦15 N natural abundance values of treatments containing non-labelled earthworms varied between different compartments, however was not significantly affected by AMF (F4,52 = 1.023, P = 0.336; earthworms: 6.39 ± 0.4‰, casts: 24.38 ± 0.7‰, roots: 1.64 ± 0.5‰, leaves: −0.62 ± 0.1‰, aphids: 0.45 ± 0.1‰). Mean ␦15 N values of earthworms, casts, roots, leaves, and aphids of treatments containing labelled earthworms differed significantly from each other (F4,52 = 116.08, P = 0.001), but were not affected by AMF (F1,55 = 0.315, P = 0.577; Fig. 1). Across AMF treatments, the highest 15 N enrichment levels were measured in casts (11,751.21 ± 1405.2‰), followed by earthworm tissue

(454.68 ± 59.4‰), roots (62.65 ± 21.8‰), leaves (50.94 ± 16.9‰) and aphids (37.75 ± 14.3‰; Fig. 1). Regarding ␦13 C, only earthworms and casts showed significantly increased values relative to −EW control treatments (Table 1). Delta 13 C values did not differ significantly between compartments (roots, leaves and aphids; F2,53 = 4.79, P = 0.543). With the exception of ␦13 C in casts (−AMF: 39.54 ± 9.9‰, +AMF: 99.21 ± 18.3‰; F1,5 = 10.13, P = 0.007) ␦ 13 C values were not significantly affected by AMF treatments. Although ␦15 N values for earthworms, casts and roots seemed to be positively affected by the presence of AMF, this was only marginally statistically significant (Fig. 1; Table 1). When considering ␦15 N of leaves and aphids as aboveground subsystem and ␦15 N of roots representing the belowground subsystem, while excluding earthworm tissue and casts as they form the vector of isotopic dispersal, a significant interaction between AMF and these subsystems was found (F1,32 = 4.61, P = 0.039). Mycorrhiza decreased 15 N signatures in the aboveground subsystem, but increased the signature in the belowground subsystem (Fig. 1). The same analysis with ␦13 C did not show a significant interaction between AMF and the subsystems (F1,32 = 0.70, P = 0.415). Neither in the 15 N nor in the 13 C signature was there a significant earthworm × AMF interaction effect (P > 0.05). Aphid ␦15 N was significantly positively correlated with leaf ␦15 N, however 15 N signals of the other compartments investigated appeared to be unrelated to each other (Table 2). Aphid ␦13 C was significantly positively correlated with leaf ␦13 C, also ␦13 C of roots and casts as well as ␦13 C of roots and leaves were significantly positively related (Table 2). The initial earthworm biomass was positively correlated with the 15 N of casts (r = 0.65, P = 0.029). Total N and C in leaves and aphids were significantly lower in +EW than in −EW treatments, but were unaffected by AMF (Table 3). At the end of the experiment, total N and C in earthworms, casts and roots were unaffected by EW and AMF treatments (Table 3).

Please cite this article in press as: Grabmaier, A., et al., Stable isotope labelling of earthworms can help deciphering belowground–aboveground interactions involving earthworms, mycorrhizal fungi, plants and aphids. Pedobiologia - J. Soil Ecol. (2014), http://dx.doi.org/10.1016/j.pedobi.2014.10.002

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Table 1 Summary of ANOVA results for the effects of earthworms (EW) or arbuscular mycorrhizal fungi (AMF) and their interactions on log ␦15 N (‰) and log ␦13 C signature in earthworm tissue, earthworm casts, roots, leaves and aphids. Significant effects (P < 0.05) are in boldface. Model degrees of freedom (df) are represented by the numerator df and the denominator df, respectively. Variable

Experimental factors Earthworms (EW) df

Log ␦15 N Earthworms Casts Roots Leaves Aphids Log ␦13 C Earthworms Casts Roots Leaves Aphids

EW × AMF

AMF

F

P

df

F

P

df

F

P

1,12 1,13 1,20 1,20 1,20

830.119 465.808 63.204 239.220 171,658

<0.001 <0.001 <0.001 <0.001 <0.001

1,12 1,13 1,20 1,20 1,20

4.144 4.134 2.443 0.973 1.935

0.064 0.063 0.134 0.336 0.179

– – 1,20 1,20 1,20

– – 0.883 0.335 1.077

– – 0.359 0.569 0.312

1,12 1,13 1,20 1,19 1,20

122.381 23.094 0.434 0.504 2.787

<0.001 <0.001 0.517 0.486 0.111

1,12 1,13 1,20 1,19 1,20

2.151 10.127 1.511 0.258 0.444

0.168 0.007 0.233 0.617 0.513

– – 1,20 1,19 1,20

– – 1.956 0.589 0.469

– – 0.177 0.452 0.501

–, missing values.

Table 2 Pearson correlation coefficients between log ␦15 N and log ␦15 C (‰) signatures of Q3 different compartments investigated in the present experiment. Only mesocosms that contained labelled earthworm were considered. Variable Log ␦15 N Casts Roots Leaves Aphids Log ␦13 C Casts Roots Leaves Aphids

Earthworms 0.352 −0.439 −0.150 −0.180 0.539 0.203 −0.137 −0.129

Casts

Roots

0.245 −0.519 −0.389

into roots, leaf tissue and phloem from where it is taken up by sap-sucking herbivores. Even across three trophic levels including detritivores, plants and herbivores the 15 N labelling signature was substantially stronger than natural abundance levels. It is concluded that a one-time addition of 15 NH4 NO3 to earthworm substrate can adequately label earthworms and associated organisms. We took most of our samples one week after introducing the labelled earthworms to mesocosms and it remains to be investigated for how long and for which sampling scale isotopic signatures are effective. A report that isotopically labelled earthworm casts lose little of their 15 N and 13 C signature even when stored in field soil for over three months (Heiner et al., 2011) suggests that trophic interactions can be studied for much longer periods than in the present study. Overall, this approach will be useful when further investigating previously unknown functional relations between above- and belowground organisms also under field conditions. Contrary to our assumption, that 13 C could be taken up by plant roots via amino acids and other organic substances (Chapin et al., 1993; Marschner, 1995; Jones et al., 2004), the amounts used in this experiment seem to be insufficient for tracing trophic interactions as 13 C enrichment in roots, leaves and aphids was in the range of natural abundance levels. An explanation for the more even distribution of 13 C tracer in different compartments could also be that 13 C cycling likely involved mineralization and uptake of 13 CO2 .

Leaves

0.237 0.125

0.781**

*

0.639 0.351 0.185

**

0.717 0.299

0.704*

Asterisks denote significant deviations from horizontal line: * P < 0.10. ** P < 0.05. *** P < 0.001.

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Discussion

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Tracking functional relationships with stable isotopes

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This study demonstrates for the first time that plants utilize nitrogen originated from earthworms and incorporate this nitrogen

Table 3 Concentrations of nitrogen and carbon (means ± SD) of earthworm tissue, earthworm casts, roots, leaves and aphids in response to earthworms (EW) or arbuscular mycorrhizal fungi (AMF). Summaries of two-way ANOVAs testing treatment effects in different compartments are mentioned on the right side of the table; significant effects are in boldbace. Model degrees of freedom (df) are represented by the numerator df and the denominator df, respectively. Different letters within each column represent significant differences (P < 0.05; n = 5–6) based on Tukey HSD posthoc mean comparisons. Variable

Treatments −EW−AMF

ANOVA results −EW+AMF

+EW−AMF

+EW+AMF

EW

Means ± SD N concentration (%) – Earthworms Casts – 1.40 ± Roots 6.18 ± Leaves 6.53 ± Aphids C concentration (%) – Earthworms – Casts 39.89 ± Roots 44.66 ± Leaves 51.91 ± Aphids

df

EW × AMF

AMF F

P

df

F

P

df

F

P

0.12a 0.14a 0.24ab

– – 1.11 ± 0.14a 6.10 ± 0.14a 6.58 ± 0.08a

11.55 0.17 1.21 5.48 5.82

± ± ± ± ±

0.45a 0.00a 0.08a 0.17b 0.18b

12.28 0.16 1.06 5.78 5.88

± ± ± ± ±

0.50a 0.00a 0.06a 0.13ab 0.18ab

1,12 1,13 1,20 1,20 1,20

3.490 8.467 1.190 12.602 15.344

0.086 0.012 0.288 0.002 0.001

1,12 1,13 1,20 1,20 1,20

1.612 0.124 4.313 0.580 0.118

0.228 0.730 0.050 0.455 0.735

– – 1,20 1,20 1,20

– – 0.458 1.782 0.001

– – 0.506 0.197 0.976

1.28a 0.65ab 0.24a

– – 41.10 ± 0.85a 43.68 ± 0.16b 51.90 ± 0.22a

44.69 5.05 38.91 41.65 51.17

± ± ± ± ±

1.27a 0.21a 0.91a 0.17cd 0.33abc

46.07 4.99 37.80 41.72 50.47

± ± ± ± ±

1.37a 0.19a 1.22a 0.31d 0.29b

1,12 1,13 1,20 1,20 1,20

0.310 0.588 0.005 0.944 3.923 0.062 42.398 <0.001 15.795 0.001

1,12 1,13 1,20 1,20 1,20

0.770 0.068 0.002 1.442 1.684

0.398 0.798 0.965 0.244 0.209

– – 1,20 1,20 1,20

– – 1.142 1.893 1.579

– – 0.298 0.184 0.223

–, missing values.

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Consistent with our expectations, AMF altered the nutrient uptake of plants resulting in an increased root 15 N signature, but a decreased leaf and aphid 15 N (significant subsystem–AMF interaction). We assume that this is because AMF facilitated nitrogen uptake by plants (Smith and Read, 2008), which resulted in a higher aboveground plant biomass and consequently diluted (decreased) 15 N levels in leaves and aphids.

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Earthworm–AMF interactions

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In the current study, effects of earthworms and interactions with AMF on growth and biomass production of the T. repens stands were not explored. The enhanced growth and biomass production of T. repens in the presence of AMF in the present study is in line with several previous studies indicating that AMF increased the growth of legumes (van der, 2002; Smith and Read, 2008; Wagg et al., 2011a, 2011b; Zaller et al., 2011a; Trouvé, 2013). Only a few studies have investigated interactions between earthworms and AMF. In the present study, root mycorrhization rates were decreased by earthworms, indicating that earthworms may have grazed on the mycelium and possibly disrupted the contact of the external hyphae from the roots. Contrastingly, some studies reported increased mycorrhization rates in the presence of earthworms (Gormsen et al., 2004; Zarea et al., 2009; Trouvé, 2013), while in several other studies no significant effects of earthworms on AMF colonization were found (Eisenhauer et al., 2009; Wurst and Rillig, 2011; Zaller et al., 2011c). Presumably, these interactions are species-specific and/or context-dependent and no general pattern can be seen. In several greenhouse experiments no interactive effects of earthworms and AMF on the performance of plants were found (Eisenhauer et al., 2009; Wurst and Rillig, 2011; Zaller et al., 2011c; Trouvé, 2013; Zaller et al., 2013b). Another study on combined effects of earthworms and AMF on plant community and diversity showed that seedling emergence and diversity was reduced by anecic earthworms in the presence of AMF (Zaller et al., 2011b). More studies on the interactions between earthworms and AMF are needed, which may allow subsequent meta-analyses and the exploration of biotic and abiotic factors shaping such belowground interactions.

422

Earthworm–AMF–aphid interactions

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Both positive and negative effects of AMF on the performance and abundance of invertebrate herbivores have been reported in previous studies (e.g. Koricheva et al., 2009; Trouvé, 2013; Zaller et al., 2013a). Earthworms have been shown to have considerable effects on nutrient cycling in soil affecting nutrient uptake and growth of plants (Brown, 1995; Curry and Schmidt, 2007; Butenschoen et al., 2009). Furthermore, earthworms can indirectly affect sap-sucking insects through altering primary and secondary metabolites and the expression of stress-responsive genes of plants (Bonkowski et al., 2001; Scheu, 2001; Blouin et al., 2005; Wurst, 2010; Trouvé, 2013). In the present experiment, AMF had no significant effects on leaf N or C concentrations and hence did not alter the performance of the aphids. Although the effect of earthworms on aphid performance was not quantitatively tested in the present study, other reports have shown that earthworms can (i) increase (Scheu et al., 1999; Wurst and Jones, 2003; Poveda et al., 2005; Eisenhauer and Scheu, 2008; Eisenhauer et al., 2010), (ii) decrease (Wurst et al., 2003; Ke and Scheu, 2008) or (iii) not affect aphid infestations (Bonkowski et al., 2001). These inconsistent results suggest that effects are context- and species-specific; combined individual effects of different soil organisms have also been reported to cancel each other out (Bradford et al., 2002; Wurst et al., 2008) or have synergistic effects on plant and herbivore performance (Eisenhauer et al., 2010).

Surprisingly, the concentrations of N and C in aphid and leaf tissue were decreased in the presence of earthworms in our experiment. This negative earthworm effect can be explained by the fact that this legume species is less responsive to soil improvements provided by earthworms through its association with N-fixing rhizobia. One other explanation may be that due to enhanced biomass production nitrogen was diluted and therefore N concentrations are only marginally affected by the amount of N bound in plant tissue. Conclusions The results of this work provide first experimental evidence that 15 N-labelled earthworms can be a powerful tool to investigate aboveground-belowground interactions across multiple trophic levels. Given the poor knowledge on how stable isotopes might be incorporated in other organisms, the next challenge would be to quantify isotopic pools and fluxes and to implement this approach in more complex, multi-species systems in the laboratory and in the field. Future research aiming at a better understanding of temporal and spatial interactions between aboveground and belowground biota might benefit from employing this methodological approach. Acknowledgments We are grateful to Norbert Schuller for help in the greenhouse and laboratory and to Wolfgang Wanek for advice on stable isotope analyses. The staff from the Department of Applied Plant Sciences and Plant Biotechnology at the University of Natural Resources and Life Sciences Vienna provided logistical support. This research was partly funded by the Austrian Science Fund (FWF project no. Q2 P20171-B16). References Arnone, J.A., Zaller, J.G., Hofer, G., Schmid, B., Körner, C., 2013. Loss of plant biodiversity eliminates stimulatory effect of elevated CO2 on earthworm casting activity in grasslands. Oecologia 171, 613–622. Bardgett, R.D., Wardle, D.A., 2003. Herbivore-mediated linkages between aboveground and belowground communities. Ecology 84, 2258–2268. Bardgett, R.D., Wardle, D.A., 2010. Aboveground–Belowground Linkages. Biotic Interactions, Ecosystem Processes, and Global Change. Oxford University Press, Oxford, UK. Blouin, M., Zuily-Fodil, Y., Pham-Thi, A.T., Laffray, D., Reversat, G., Pando, A., Tondoh, J., Lavelle, P., 2005. Belowground organism activities affect plant aboveground phenotype, inducing plant tolerance to parasites. Ecol. Lett. 8, 202–208. Bohlen, P.J., Parmelee, R.W., McCartney, D.A., Edwards, C.A., 1997. Earthworm effects on carbon and nitrogen dynamics of surface litter in corn agroecosystems. Ecol. Appl. 7, 1341–1349. Bonkowski, M., Geoghegan, I.E., Birch, A.N.E., Griffiths, B.S., 2001. Effects of soil decomposer invertebrates (protozoa and earthworms) on an above-ground phytophagous insect (cereal aphid), mediated through changes in the host plant. Oikos 95, 441–450. Bonkowski, M., Griffiths, B.S., Ritz, K., 2000. Food preferences of earthworms for soil fungi. Pedobiologia 44, 666–676. Bradford, M.A., Jones, T.H., Bardgett, R.D., Black, H.I.J., Boag, B., Bonkowski, M., Cook, R., Eggers, T., Gange, A.C., Grayston, S.J., Kandeler, E., McCaig, A.E., Newington, J.E., Prosser, J.I., Setälä, H., Staddon, P.L., Tordoff, G.M., Tscherko, D., Lawton, J.H., 2002. Impacts of soil faunal community composition on model grassland ecosystems. Science 298, 615–618. Brown, G.G., 1995. How do earthworms affect microfloral and faunal community diversity? Plant Soil 170, 209–231. Brussaard, L., 1999. On the mechanisms of interactions between earthworms and plants. Pedobiologia 43, 880–885. Butenschoen, O., Marhan, S., Langel, R., Scheu, S., 2009. Carbon and nitrogen mobilisation by earthworms of different functional groups as affected by soil sand content. Pedobiologia 52, 263–272. Chapin, F.S., Moilanen, L., Kielland, K., 1993. Preferential use of organic nitrogen for growth by a nonmycorrhizal arctic sedge. Nature 361, 150–153. Cooke, A., 1983. The effects of fungi on food selection by Lumbricus terrestris (L.). In: Satchell, J.E. (Ed.), Earthworm Ecology. From Darwin to Vermiculture. Chapman and Hall, London, pp. 365–374. Coûteaux, M.-M., Bottner, P., 1994. Biological interactions between fauna and the microbial community in soils. In: Ritz, K., Dighton, J., Giller, K.E. (Eds.), Beyond the Biomass. Wiley-Sayce, London, pp. 159–172.

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