Effects of silver nanoparticles on soil microorganisms and maize biomass are linked in the rhizosphere

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Effects of silver nanoparticles on soil microorganisms and maize biomass are linked in the rhizosphere Wouter Sillen a, Sofie Thijs a, Gennaro Roberto Abbamondi a, b, Jolien Janssen a, Nele Weyens a, Jason White c, Jaco Vangronsveld a, * a b c

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Hasselt University, Center for Environmental Sciences, Agoralaan Building D, 3590 Diepenbeek, Belgium National Research Council of Italy, Institute of Biomolecular Chemistry, Via Campi Flegrei 34, Pozzuoli (Napoli) 80078, Italy Connecticut Agricultural Experiment Station, Department Analytical Chemistry, 123 Huntington Street, New Haven, CT, USA

a r t i c l e i n f o

a b s t r a c t

Article history: Received 11 April 2015 Received in revised form 13 August 2015 Accepted 14 August 2015 Available online xxx

Silver nanoparticles hold great promise as effective anti-microbial compounds in a myriad of applications but may also pose a threat to non-target bacteria and fungi in the environment. Because microorganisms are involved in extensive interactions with many other organisms, these partner species are also prone to indirect negative effects from silver nanoparticles. Here, we focus on the effects of nanosilver exposure in the rhizosphere. Specifically, we evaluate the effect of 100 mg kg
1 silver nanoparticles on maize plants, as well as on the bacteria and fungi in the plant's rhizosphere and the surrounding bulk soil. Maize biomass measurements, microbial community fingerprints, an indicator of microbial enzymatic activity, and carbon use diversity profiles are used. Hereby, it is shown that 100 mg kg
1 silver nanoparticles in soil increases maize biomass, and that this effect coincides with significant alterations of the bacterial communities in the rhizosphere. The bacterial community in nanosilver exposed rhizosphere shows less enzymatic activity and significantly altered carbon use and community composition profiles. Fungal communities are less affected by silver nanoparticles, as their composition is only slightly modified by nanosilver exposure. In addition, the microbial changes noted in the rhizosphere were significantly different from those noted in the bulk soil, indicated by different nanosilver-induced alterations of carbon use and community composition profiles in bulk and rhizosphere soil. Overall, microorganisms in the rhizosphere seem to play an important role when evaluating the fate and effects of silver nanoparticle exposure in soil, and not only is the nanosilver response different for bacteria and fungi, but also for bulk and rhizosphere soil. Consequently, assessment of microbial populations should be considered an essential parameter when investigating the impacts of nanoparticle exposure. © 2015 Elsevier Ltd. All rights reserved.

Keywords: Maize Bacteria Fungi Rhizosphere Silver nanoparticles ARISA

1. Introduction Microorganisms form significant interactions with a wide array of biota in terrestrial ecosystems, often with important consequences for all species present. Terrestrial plant species are colonized by a large number of microorganisms in several morphological regions, including interior tissues (endophytes), on their leaves (epiphytes) and in their rhizosphere. In the rhizosphere, the microbial genes greatly outnumber the plant genes (Mendes et al., 2013), resulting in a microbiome that has been referred to

* Corresponding author. Tel.: þ32 11 268331; fax: þ32 11 268399. E-mail address: [email protected] (J. Vangronsveld).

as the plant's second genome. Just like the human gut microbiome, the plant's rhizosphere microbiome has been implicated in many functions that support its host's well-being. For example, rhizosphere microorganisms can increase nutrient uptake, protect against pathogens, enhance abiotic stress tolerance and promote the development of the plant (Berendsen et al., 2012). However, some rhizosphere organisms are also capable of causing plant disease or impeding plant growth. Alternatively, plants also influence the microbial community within their rhizosphere, largely mediated through root structure and exudates that determine the physical and chemical conditions within the plant root zone. As root structure and exudates change during the plant's life cycle, the conditions in the rhizosphere are altered, likely selecting for a

http://dx.doi.org/10.1016/j.soilbio.2015.08.019 0038-0717/© 2015 Elsevier Ltd. All rights reserved.

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different microbial community. Other factors such as xenobiotics can disturb the rhizosphere conditions, having both direct and indirect consequences for the plant and its microbial community. The dramatic increase in the use of silver nanoparticles (AgNP) has resulted many beneficial applications but also raised some concern. On the one hand, nanotechnology has begun to make efficient use of the well-known anti-microbial properties of silver. On the other hand, the increasing quantities in which AgNP are produced and applied, as well as the unique characteristics resulting from the nanoscale size, have led to concern over the toxicological implications of exposure for nontarget species. Results from studies with several plant species are largely in agreement over the phytotoxic effects of AgNP in artificial conditions such as hydroponic systems. AgNP have been demonstrated in hydroponics to impair root elongation, seed germination and plant biomass production of plants like zucchini, cucumber, onion, rye grass and rice (Stampoulis et al., 2009; Anjum et al., 2013; Dimkpa, 2014; Gardea-Torresdey et al., 2014). However, in the much scarcer soilbased studies, the concentrations at which a negative effect of AgNP on plants are observed are often higher than in hydroponics studies, and the proportion of reported neutral AgNP effects on plant growth increases (Dimkpa, 2014). As mentioned, the interest in AgNP mainly originates from its well-known potency as a broad spectrum antimicrobial agent effective against diverse species of Gram-positive and Gram-negative bacteria (Morones et al., 2005; Kim et al., 2007), as well as against several fungal species, although reports of this anti-fungal activity are considerably less numerous (Panacek et al., 2009). As both bacteria and fungi are present in the rhizosphere of plants, the antimicrobial effects from AgNP exposure, may have significant implications for the host plant health. Notably, the amount of research investigating the implications on rhizosphere-based nanoparticle exposure on both the microbes present and the supporting host plant is scarce. This study investigates the effects of AgNP exposure on maize plants and the bacteria and fungi in the plant rhizosphere. The strongly microorganism-oriented toxicity of AgNP in soil will be used to increase our understanding on the plantemicrobe interaction in the rhizosphere, in particular under abiotic stress such as from NP exposure. Maize is chosen as the experimental crop because of its importance in global agriculture, and because of the likelihood of significant AgNP-exposure as AgNP may enter the agro-ecosystem through several pathways: during manufacturing, from use of nano-enabled agrichemical products, and from the application of NP-containing biosolids. Specifically, maize plants are grown for 75 days in soil containing 100 mg kg
1 AgNP, and the effects of this exposure on biomass production and on the activity and community structure of its associated bacteria and fungi are monitored over time. This concentration of 100 mg kg
1 AgNP is high when compared to data of studies that examine the fate of unintentionally released AgNP from non-agricultural related products (Gottschalk et al., 2009; Simonin and Richaume, 2015). However, because agricultural maize fields accumulate also AgNP from biosolids and nano-enabled agrichemical sources, AgNP concentrations are higher in these systems than in those that just suffer from unintentional release. Using a concentration like the one we applied will provide heretofore unavailable data of the effects of a new contaminant at a concentration on the doseeresponse curve where an effect is anticipated.

Houston, Texas, USA. The experimental soil was collected from the top 30 cm of an agricultural corn field in Diepenbeek, in the eastern part of Belgium (50 560 05.300 N 5 240 41.200 E), and was characterized as sandy loam (55% sand, 30% silt, 15% clay) with a pH of 6.98, an electrical conductivity (EC) of 335 mS cm
1 and an effective cationexchange capacity (CEC) of 20.7 meq/100 g. After collection, the soil was 6 mm sieved and homogenized. Zea mays variety LG 30.223 seeds were obtained from LimaGrain Belgium. In order to evaluate the effects of AgNP-exposure on plants and microbial communities both in bulk soil and rhizosphere over time, the following experimental design was applied. A total of 120 pots with 1 kg of soil each was used, with half of them being amended with 100 mg kg
1 AgNP by mechanical mixing during 5 min. As mentioned above, we recognize that this exposure concentration is high (Simonin and Richaume, 2015), although such concentrations could be achieved in the future on maize fields due to the fact that they accumulate nanoparticles from multiple sources (biosolids, nano-enabled agrichemicals). Moreover, we used this high level to establish a baseline response in the system. The other half of the pots, without AgNP, were used as controls. Half of both control and AgNP pots were planted with a maize seed; the other half of the pots remained unplanted. Before planting, maize seeds were soaked in tap water overnight. All pots were randomly placed in a greenhouse under the following conditions: photoperiod of 14 h daylight, a temperature cycle of 22 C/18  C and a relative humidity of 60%. After 16, 25, 39, 53 and 75 days, plants were harvested and samples were taken from the rhizosphere, here defined as soil that remained attached to the root after light shaking, and the bulk soil for microbial community analysis. Specifically, at each harvesting point, 6 pots with maize plants (3 control and 3 AgNP-exposed), and 6 pots with bare soil (3 control and 3 AgNP-exposed) were randomly selected; the former 6 being used for plant measurements and rhizosphere samples, and the latter 6 for bulk soil samples. The total leaf length of each harvested plant was determined, and harvested shoots and roots were oven dried at 60  C and subsequently weighed. Bulk soil and rhizosphere samples were used for fluorescein diacetate (FDA) hydrolysis analysis, the Biolog EcoPlate™ assay and bacterial and fungal automated ribosomal intergenic spacer analysis (ARISA) fingerprinting. 2.2. Biolog EcoPlate™ analyses Samples from rhizosphere and bulk soils were used immediately after harvest for community-level physiological profiling (CLPP) with Biolog EcoPlates™ (Biolog, California, USA). For each sample, 1 g of soil was dissolved in 10 ml 0.01 M phosphate buffer (pH ¼ 7.2), and after thorough shaking, the solution was further diluted 1:20. After settling of remaining particles, 120 ml was dispensed into each well of all 31 carbon sources (3 replicates per plate). The plates were incubated in the dark at 25  C, and absorbance at 590 nm of all wells per plate was measured twice every 24 h using a FLUOstar® Omega plate reader, with the final measurement taking place after 94 h. Absorbance data were used to calculate the average well color density (AWCD), richness (R) with threshold equal to 0.25, and Shannon-Weaver index (H) per treatment. Absorbance values per carbon source were also used separately in subsequent statistical analyses. 2.3. FDA assay

2. Material and methods 2.1. Material and experimental set-up Uncoated silver nanoparticles (99.99% purity, 20 nm diameter) were acquired in solid form from US Research Nanomaterials, Inc.,

Immediately after harvesting, samples from bulk and rhizosphere soils were used for the FDA-hydrolysis assay (Schnurer and Rosswall, 1982; Adam and Duncan, 2001). Briefly, 1 g of soil (replicated twice) for each of the 3 samples per treatment were incubated for 3 h at 30  C with 50 ml of 60 mM sodium phosphate

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buffer (pH ¼ 7.2) and 0.5 ml of a FDA stock solution (2 mg fluorescein diacetate ml
1). In order to stop the reaction, 2 ml of acetone was added, and the resulting mixture was centrifuged. The absorbance at 490 nm of the filtered (Whatman No2) supernatant was measured. 2.4. ARISA fingerprinting Total DNA was extracted from the harvested bulk and rhizosphere soils using the PowerSoil® DNA Isolation Kit (Mo Bio Laboratories Inc., California, USA). Bacterial 16Se23S ITS DNA was amplified using the primer pair S-D-Bact-1522-b-S-20 (eubacterial rRNA small subunit, 50 -TGCGGCTGGATCCCCTCCTT-30 ) and L-D-bact-132-a-A-18 (eubacterial rRNA large subunit, 50 CCGGGTTTCCCCATTCGG-30 ) (Normand et al., 1996). Fungal ITS15.8S-ITS2 DNA was amplified with the primer pair 2234C (at 30 end of 18S gene, 50 -GTTTCCGTAGGTGAACCTGC-30 ) and 3126T (at 50 end of 28S gene, 50 -ATATGCTTAAGTTCAGCGGGT-30 ) (Sequerra et al., 1997). PCR amplification was carried out in 50 ml reaction mixtures containing 15e40 ng ml
1 of environmental DNA, 1  PCR buffer, 1 U of Platinum® High Fidelity Taq DNA polymerase (Invitrogen, California, USA), 0.2 mM (each) dNTP, 2 mM MgSO4 and 0.2 mM (each) primer. Cycling conditions for the bacterial primer pair consisted of a hot start at 94  C for 3 min and 25 subsequent cycles consisting of 94  C for 1 min, annealing at 55  C for 30 s and elongation at 72  C for 1 min. Finally, an elongation step at 72  C for 5 min was followed by cooling at 4  C. The reaction mixture with the fungal primers was held at 94  C for 3 min, followed by 30 cycles of 94  C for 45 s, 57.5  C for 1 min, 72  C for 2 min, and a final extension at 72  C for 7 min. Amplified reaction mixtures were loaded onto DNA-1000chips that were prepared according to the manufacturer's recommendations, allowing the product fragments to be resolved with an Agilent 2100 Bioanalyzer (Agilent Technologies, USA). The 2100 Expert Software (Agilent Technologies) was used to digitalize the ARISA fingerprints, resulting in electrophoregrams in ASCII formats that were processed using the StatFingerprints package (Michelland et al., 2009), in the 2.13.0 version of the R project (The R Foundation for Statistical Computing, Vienna, Austria). 2.5. Statistical analyses All statistical analyses were performed using the 3.1.1 version of R (The R Foundation for Statistical Computing, Vienna, Austria). ANOVA or the KruskaleWallis rank sum test were used when checking for significant differences in parameter values, depending on whether or not, respectively, normality and homoscedasticity assumptions were met. Depending on the same assumptions, two by two comparisons were made using either Tukey's honest significant differences tests or pairwise Wilcoxon rank sum tests. Multivariate datasets were analyzed using linear discriminant analysis (LDA) for the CLPP data, non-metric multidimensional scaling (NMDS) with the BrayeCurtis distance metric for the ARISA fingerprints, and canonical correspondence analysis (CCA) for an integration of all responses. In addition, analyses of similarities (ANOSIM) were performed when checking for significant differences between groups with multivariate data, and Mantel tests were used to test for correlations between distance matrices (BrayeCurtis for fingerprint data, Euclidean in the other cases). 3. Results 3.1. Plant response For all measured parameters, there was a statistically significant difference between the AgNP-exposed and the non-exposed plants

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at some harvesting point. As apparent from Fig. 1, the dry biomass of the aboveground and belowground AgNP-exposed plant parts was significantly higher than that of the non-exposed plants after 75 days of growth (p < 0.1 and p < 0.05, respectively). At other time points there was a distinguishable trend of biomass increase with AgNP-exposure. At no stage was a trend of decreased biomass with AgNP-exposure observed. Total leaf length displayed a pattern similar to the dry biomass, but showed an additional significant (p < 0.05) increase after 39 days of growth under AgNP-exposure. The size of the soil containers that were used to grow the plants seemed to have an effect that was mainly noticeable at the harvesting point after 75 days, as plants at this point displayed a trend (not statistically significant) of decrease in dry root biomass and total leaf length, compared to the earlier harvesting point at 53 days. 3.2. Microbial activity and metabolic diversity As indicated by the FDA-assay (Fig. 2), microbial activity decreased after the second harvesting point, but remained generally stable afterward, not considering the different treatments. Due to technical reasons, the FDA-assay was not performed at the first sampling point after 16 days of growth. When statistically significant differences were found between treatments at some time point, these differences always pointed out that FDA-hydrolysis activity in the non-exposed rhizosphere was higher than at least one of the other three sample types. After 39 days of growth, the non-exposed rhizosphere activity was higher than the activity in the AgNP-exposed and non-exposed bulk soil, and in the AgNPexposed rhizosphere (p < 0.01). After 75 days of growth, the activity in the non-exposed rhizosphere was higher than the activity in the AgNP-exposed rhizosphere and bulk soil (p < 0.1 and p < 0.05, respectively). In order to create the CLPP, the Biolog EcoPlates™ were incubated for 94 h, and Abs590nm values after this incubation period were used to calculate the average well color density (AWCD), the richness (R) and the Shannon-Weaver index (H) of all sample types. In Fig. 3, only the results for AWCD are shown, as the results for the other two parameters displayed nearly identical relative proportions and statistical significances as this parameter. When differences in AWCD were found, AgNP-exposed samples always had a lower value than non-exposed samples. The samples after 16 and 53 days indicated that values of AgNP-exposed bulk soil were significantly lower than those of the non-exposed (p < 0.01), while AgNP-exposed rhizosphere showed values similar to the nonexposed. At the sampling points after 39 and 75 days, however, both AgNP-exposed bulk soil and rhizosphere had lower values than the non-exposed samples (p < 0.01 and p < 0.05, respectively). In addition to the values after 94 h of incubation, the change in AWCD during the incubation period allows for discrimination between the different sample types (Fig. 4). The AgNP-exposed samples showed a less steep value increase and often a longer lag-period than the non-exposed samples, with the bulk soil samples showing the biggest difference whereas the rhizosphere samples resemble more the non-exposed samples. When considering the CLPP results of all carbon sources individually, more detailed distinctions can be made. Fig. 5A shows a linear discriminant analysis (LDA) of the Abs590nm values after 94 h of incubation, using the data of harvesting at 16 and 53 days. Additionally, group means for all sample types are depicted with a probability ellipse (standard deviation, p ¼ 0.68), and the font size of the carbon sources corresponds to the score of each source on a SIMPER analysis distinguishing between AgNP-exposed and nonexposed bulk soil samples at these harvesting points. Similarly to the results in Fig. 3, the findings demonstrate that at these

Please cite this article in press as: Sillen, W., et al., Effects of silver nanoparticles on soil microorganisms and maize biomass are linked in the rhizosphere, Soil Biology & Biochemistry (2015), http://dx.doi.org/10.1016/j.soilbio.2015.08.019

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harvesting points, there was a clear AgNP effect on the carbon use pattern, and that bulk and rhizosphere soil were very similar in their pattern in the non-exposed samples, but very different in the AgNP-exposed samples. The carbon use pattern of the AgNP-exposed rhizosphere was more similar to that of the nonexposed samples, than was the carbon use pattern of the AgNPexposed bulk soil. Fig. 5B shows a linear discriminant analysis of the Abs590nm values after 94 h of incubation, using the data of

harvesting points 39 and 75 days. Group means for all sample types are depicted with a probability ellipse (standard deviation, p ¼ 0.68), and the font size of the carbon sources corresponds to the score of each source on a SIMPER analysis distinguishing between AgNP-exposed and non-exposed rhizosphere samples of these harvesting points. This figure shows that, similarly to the pattern at harvesting points 16 and 53 days, there was a clear general AgNP effect and that bulk soil and rhizosphere were more similar to each

Please cite this article in press as: Sillen, W., et al., Effects of silver nanoparticles on soil microorganisms and maize biomass are linked in the rhizosphere, Soil Biology & Biochemistry (2015), http://dx.doi.org/10.1016/j.soilbio.2015.08.019

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Differences between sample types were also analyzed by applying Analysis of Similarity (ANOSIM) to the CLPP data. In the bulk soil, a change in carbon use pattern as AgNP-response was observable at harvesting points 16, 39, 53 and 75 days, while in the rhizosphere, a change was noticeable only at harvesting points 39 and 75 days (R > 0.75). Bulk soil and rhizosphere never had a different carbon use pattern in the non-exposed samples, while in the AgNP-exposed samples the two soil types differed at harvesting points 39, 53 and 75 days (R > 0.75).

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other in the non-exposed samples than in the AgNP-exposed samples. However, at these harvesting points, the carbon use pattern of AgNP-exposed rhizosphere was more different from that of the non-exposed rhizosphere than was the case at harvesting points 16 and 53 days. Nonetheless, this difference did not result from a similar change in carbon use pattern as in the bulk soil, as indicated by the SIMPER results in Fig. 5A and B. While C-sources like D-xylose, phenylethylamine and 4-hydroxy benzoic acid seemed to contribute the most to the difference between nonexposed and AgNP-exposed bulk soil at harvesting points 16 and 53 days, it was mainly glycogen, alpha-cyclodextrin and itaconic acid that distinguished non-exposed and AgNP-exposed rhizosphere at 39 and 75 days. The general pattern shown by the LDA's pointed out that no C-sources were strictly typical for any sample type in general, and that differences were mainly caused by generally lower C-oxidation values in AgNP-exposed samples, compared to non-exposed samples.

Bacterial fingerprints through ARISA were made of the DNA extracted from bulk and rhizosphere soils at all sampling points. Processed fingerprints were used in an NMDS-analysis with the BrayeCurtis distance metric (Fig. 5C, stress ¼ 0.088), with a probability ellipse (standard deviation, p ¼ 0.68) for each sample type. AgNP-exposure in general affected the bacterial community both in the bulk and rhizosphere soil. In the bulk soil, the effect appeared at 16 days, while in the rhizosphere, the effect was observed after 25 days (ANOSIM R > 0.75). A significant difference between bulk and rhizosphere soil communities was not observed at any harvesting point for non-exposed samples, but there was such a difference for the AgNP-exposed samples at harvesting points 39 and 75 days (R > 0.6). From the four different sample types, AgNP-exposed rhizosphere showed the greatest inner variation. At harvesting points 16, 25 and 53 days, these samples resembled each other, while the community structures at harvesting points 39 and 75 days differed strongly from the average of this group, as well as from each other. 3.4. Fungal community structure Soil DNA extracted from bulk soil and rhizosphere at all harvesting points was used to create ARISA fungal fingerprints. In Fig. 5D, the result of an NMDS-analysis with the BrayeCurtis distance metric of these processed fingerprints is shown (stress ¼ 0.18). As with bacterial communities, there was a general

Please cite this article in press as: Sillen, W., et al., Effects of silver nanoparticles on soil microorganisms and maize biomass are linked in the rhizosphere, Soil Biology & Biochemistry (2015), http://dx.doi.org/10.1016/j.soilbio.2015.08.019

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Fig. 5. Multivariate statistical analyses of the CLPP and ARISA fingerprint data. A, B: linear discriminant analysis (LDA) of the CLPP Abs590nm values after 94 h of incubation of control and AgNP-exposed samples of rhizosphere and bulk soil at the harvesting points (A) 16 and 53 days, or (B) 39 and 75 days. Group means for the sample types are shown with a probability ellipse (standard deviation, p ¼ 0.68). The font size of the carbon sources depicts the score of each source on a SIMPER analysis that distinguishes between AgNP-exposed and non-exposed bulk soil samples of the respective harvesting points. C, D: NMDS-analysis with the BrayeCurtis distance metric of the (C) bacterial (stress ¼ 0.088) and (D) fungal (stress ¼ 0.18) ARISA fingerprints of control and AgNP-exposed samples of rhizosphere and bulk soil, with the harvesting point (in days) mentioned next to each sample. Probability ellipses (standard deviation, p ¼ 0.68) are shown for each sample type.

effect of AgNP-exposure on fungal communities in both bulk and rhizosphere soils, although the effects were less pronounced than observed with the bacterial communities, and often were not of statistical significance. The AgNP exposure only had a significant effect in the bulk soil at 25 days, and in the rhizosphere, at 25 and 53 days (ANOSIM R > 0.75). There were no significant differences between bulk soil and rhizosphere fungal communities at any harvesting point in the AgNP-exposed samples and non-exposed soils. The very low variability that existed among the AgNP-exposed bulk soil samples at the different harvesting points is noteworthy when compared to the variability in all other three samples types. 3.5. Correspondence between plant and microorganism AgNPresponses Mantel tests with 1000 permutations were performed on the distance matrices of the plant parameters (Euclidean distance), CLPP carbon use data (Euclidean distance), bacterial ARISA fingerprints (BrayeCurtis distance) and fungal ARISA fingerprints (BrayeCurtis distance), with the data of all harvesting points. The bacterial and fungal fingerprints had a significant correlation coefficient (r) of 0.31 (p < 0.05; H0: r  0), but these two datasets differed strongly in their respective correlations with the plant and CLPP data. Namely, while the bacterial fingerprints data had rvalues of 0.2 (p < 0.05; H0: r  0) and 0.13 (p < 0.1; H0: r  0) with

the CLPP and plant data, respectively, the respective values for the fungal fingerprint data were the non-significant 0.06 and 0.02 (p > 0.2; H0: r  0). In order to further explore the relationship between the measured responses, a canonical correspondence analysis (CCA) was performed on the bacterial and fungal ARISA fingerprints from the rhizosphere, constrained by the plant parameters, FDAhydrolysis activity and CLPP parameters of the corresponding samples (Fig. 6A & B). Similarly to Fig. 5C, the CCA with the bacterial fingerprints shows a clear separation between the fingerprints of AgNP-exposed and non-exposed samples. In addition, it is shown that the plant parameters were positively correlated with this AgNP-effect, while bacterial/microbial metabolic parameters were negatively correlated with the AgNP-effect. The CCA with the fungal fingerprints shows, similarly to Fig. 5D, an AgNP-effect that is less strong than that observed with the bacterial fingerprints. In addition, the correlation of the AgNP-effect with FDA-hydrolysis and, especially, the plant parameters was far less pronounced. 4. Discussion 4.1. Effect of AgNP on microbial communities Because of silver's well-documented toxicity to microbes, a significant effect of AgNP-exposure on both bacteria and fungi

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should be expected. Furthermore, since fungi are known to cope with metal-stress more effectively than bacteria, the latter should display the greatest impact of AgNP exposure. The results documented here do indeed confirm these considerations. Also, there appears to be an effect of incubation time, as statistically significant differences appear for the most part at the later harvesting points. Total microbial activity, indicated by the FDA-assay, showed statistically significant differences between some sample types only at the 39 and 75 day harvesting points (Fig. 2). These differences consistently demonstrate lower activity in AgNP-exposed samples compared to non-exposed samples. FDA can be hydrolyzed by esterases, proteases and lipases in soil; these are enzymes that are widespread in soil, especially among the major decomposers, bacteria and fungi (Schnurer and Rosswall, 1982). These enzymatic activities are known to be impaired when the microbial community is exposed to stress; in fact, silver nanoparticles already have previously been found to decrease FDAhydrolysis in soil. However, greater differences are found when looking at specific soil exoenzymes individually (Shin et al., 2012). The FDA-assay produces a general picture of enzymatic activity, but lacks resolution because many bacteria and fungi possess the FDA-hydrolysis capacity. Also, changes in the microbial community can occur without being reflected in the FDA-assay, since some soil community members have a more rapid metabolism than others. Changes in the bacterial community under AgNP-exposure are strongly pronounced on both a physiological and phylogenetic level. Nanosilver exerted strong toxicity on bacteria, with the general effects visible throughout all harvesting points as determined by the ARISA fingerprinting and CLPP results (Fig. 5). However, the magnitude of the effect varies between sample types and harvesting points, with more statistically significant differences occurring at later harvesting points. This effect of incubation time could be attributed to the specific dissolution rate of AgNP in the complex soil matrix, and to the effect of the plant growth stage on the bacterial community (Gomes et al., 2001; Baudoin et al., 2002;

Anjum et al., 2013). Nanosilver's toxicity to bacteria is welldocumented and investigated in artificial environments and also appears to play a similarly important role in the more complex soil environment. A few studies have highlighted this effect before, by investigating the effects of AgNP exposure on the bacterial community in sludge (Sun et al., 2013; Yang et al., 2014) and soil (Hansch and Emmerling, 2010; Kumar et al., 2011). The slower increase in AWCD of AgNP-exposed samples throughout all harvesting points (Fig. 4) indicates that already from the early periods of exposure, AgNP cause a decrease in the size (and possibly in the metabolic activity and substrate use efficiency) of the bacterial community. In addition, the differences in physiological diversity, as indicated by the carbon use patterns in the CLPP, arise just as soon in the bulk soil. The community ARISA profiles are altered by AgNP at the early harvesting points, but the effect becomes more pronounced with time. Interestingly, there are important differences between bulk and rhizosphere soils that will be discussed below. Similarly to the bacterial community, the fungal community shows structural AgNP-responses, although the changes are generally less pronounced than for the former. Shifts in the fungal community can be seen through the ARISA fingerprints (Fig. 5D), but these shifts are not as strong and of less statistical significance than the shifts in the bacterial community. This difference can be related to the general observation that eukaryotic and prokaryotic microorganisms display different sensitivity to silver (Pshennikova et al., 2011), with fungi being more tolerant to silver (Kathiresan et al., 2010) and heavy metals in general (Hiroki, 1992). These different bacterial and fungal responses have also been reported by others (Gryndler et al., 2012; Kumar et al., 2014). Remarkably, our results show a small variance among the AgNP-exposed bulk soil samples of the different harvesting points, when compared to the variation of the other three sample types. Apparently, AgNP-exposure in bulk soil leads to a confined fungal community structure that is stable over time, unlike the rhizosphere soil that hosts a community displaying different properties.

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4.2. The rhizosphere-effect and its implications for the AgNP-effect on microbial communities In the rhizosphere, the living conditions of microorganisms are substantially different from those in the bulk soil. Roots and their exudates physically and chemically alter the microenvironment, resulting in a microbial community that is more numerous but less diverse than in bulk soil (Roesch et al., 2007; Weinert et al., 2011). Moreover, these specific conditions in the rhizosphere can also change the properties of AgNP, resulting in changes in their toxicity to microorganisms (Dimkpa, 2014). The presence of a higher number of microorganisms increases the number of microbial factors such as extracellular polymeric substances that are capable of reducing AgNP-toxicity (Joshi et al., 2012; Karunakaran et al., 2013). Conclusively, the rhizosphere is expected to harbor a different microbial community than does the bulk soil (rhizosphere effect), and its community is expected to respond differently to AgNP-exposure. In our study, results indicated that bulk and rhizosphere soil microbial communities differed only to a limited extend in activity, metabolic diversity and community structure. The FDA-assay (Fig. 2) indicates that at two harvesting points, the total microbial activity in the AgNP-exposed rhizosphere was lower than in the non-exposed rhizosphere. This, however, was the only retrievable statistically significant indication of the rhizosphere effect on the microbial community. Fingerprinting techniques and CLPP as they were used here, appear to be unable to pick up further differences. Far more obvious are the differences in AgNP-response between rhizosphere and bulk soil. AgNP-exposure makes the carbon use profile of the bulk soil shift further away from the control than it does for the rhizosphere (Figs. 3 and 5A and B). Also, at the harvesting points where the carbon use profile of the rhizosphere community is significantly altered, these AgNPinduced changes in the rhizosphere are related to different carbon sources than is the case for the AgNP-induced changes in the bulk soil (Fig. 5A and B). The bacterial and fungal ARISA fingerprints indicate that the AgNP-exposure produces community structure alterations that are more variable in the rhizosphere than in the bulk soil (Fig. 5C and D). Also, it takes longer before AgNP-exposure has a significant effect on the bacterial community structure in the rhizosphere than it does in the bulk soil, and at two harvesting points, the AgNP-exposed bulk soil and rhizosphere bacterial communities differ significantly. As such, it can be concluded that the rhizosphere microbial community is less sensitive to AgNPexposure than the bulk soil, and organisms within the plant root zone respond differently to AgNP-exposure than does the bulk soil microbial community. 4.3. AgNP-effects on maize and on maize microbial community are linked The evaluation of plant parameters has revealed that the AgNPexposure conditions used in our experiments did not induce observable negative effects on maize biomass, and at two of the later harvesting points, the nanoparticles significantly increased plant mass. This may be surprising since AgNP are frequently cited as phytotoxic nanoparticles; a number of studies in hydroponic systems with diverse plant species have indicated negative effects of AgNP on plant growth or other parameters (Stampoulis et al., 2009; Miralles et al., 2012; Gardea-Torresdey et al., 2014). In soilbased conditions, however, the complex array of biotic and abiotic processes can serve to significantly modify the overall toxicity profile (Anjum et al., 2013; Dimkpa, 2014). For example, the interaction of AgNP with the soil environment is known to change particle physical and chemical properties which subsequently can then alter NP stability, transport, availability, and subsequent

toxicity to biota (Anjum et al., 2013). Our results highlight another factor that might account for the lack of AgNP toxicity to maize in soil: the interaction with microorganisms in the rhizosphere. Our results indicate that the effects of AgNP-exposure on maize and on the bacterial community in its rhizosphere are negatively correlated. After 75 days, maize plants showed significantly higher dry biomass production when they were exposed to AgNP. Additionally, after both 39 and 75 days, maize total leaf length was significantly higher under AgNP-exposure. Notably, at those two harvesting points, the bacterial rhizosphere community exhibited significant changes under AgNP-exposure. Carbon use diversity decreased and shifted, and the community structure, as determined by DNA fingerprinting, changed significantly. These changes are also apparent from the FDA-assay, showing that the total microbial activity was significantly lowered by AgNP-exposure only at these two harvesting points. This decrease in activity indicates that the changes in the microbial community are caused by the negative effects of AgNP on bacteria, which in general have a higher activity level than fungi. The correlation between bacterial community and plant responses is visible in Fig. 6A, as the AgNP-related shift in community fingerprint as well as the bacterial metabolic parameters are negatively correlated with the plant parameters. The AgNP-response of the fungal community structure, however, shows almost no correlation to the plant AgNP-response. This can be seen in Fig. 6B, especially when compared to Fig. 6A, as the general AgNP-effect on the fungal community is less clear and not as well correlated to the plant AgNP-response as the bacterial community. The discrepancy between bacteria and fungi in plant AgNP-response correlation is further demonstrated by the Mantel test results. Interestingly, however, the harvesting points at which the fungal rhizosphere community structure was significantly altered by AgNP-exposure, namely the harvesting point after 25 and 53 days, are points where bacterial community structure and plant parameters showed no AgNP-influence. Alterations in fungal rhizosphere community structure therefore do not coincide with improved maize biomass production under AgNP-exposure. Strikingly, although the AgNP-response of the bacterial but not the fungal community is correlated to the maize AgNP-response, the bacterial and fungal community structure changes overall in bulk and rhizosphere soil are positively correlated. This highlights the numerous interactions between these two groups of organisms in soil that influence each other's community structures and their response to environmental stresses such as AgNP-exposure. These interactions might also be responsible for the relation between the negative effect of AgNP exposure on bacteria and the positive effect on maize growth, as fungi and their positive effects on maize growth could proliferate in an AgNP-affected bacterial community. This possible link is a topic of ongoing investigation. The extensive interaction platform between microbes and plants that exists in the rhizosphere, is already known to have farreaching implications for both partners. Our results show a maize biomass increase in case of AgNP-exposure, but only when also the bacterial community in its rhizosphere undergoes significant alterations and the fungal community remains largely unchanged. The rhizosphere microbial community apparently is altered in a way that benefits maize growth. This can originate from a diverse array of either increased plant-beneficial characteristics or decreased plant-disadvantageous properties in the rhizosphere microbial community, mainly mediated by shifts in the bacterial community. There has been significant focus on AgNP in phytopathogens management research (Servin et al., 2015), and phytopathogen suppression is also a possible explanation for the AgNPinduced increase in plant growth. However, other microorganismmediated processes such as fungal proliferation, suggested above, and increased mycorrhization are equally likely to explain the

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observed effects. Additional research is needed to unambiguously state the mechanism behind the AgNP-induced plant growth increase. Our results highlight the importance of rhizosphere interactions and additionally pose that these interactions have important consequences for the impact of foreign substances such as AgNP on multiple members of the system. Several kinds of NPs have already been shown to directly affect plant growth and productivity in artificial systems, but the effects of NPs on rootassociated microbes are very likely to play an important role in actual terrestrial systems. Clearly, additional investigations addressing the impacts of widely used materials such as AgNP should occur under soil-based conditions and the impact of exposure on plant-associated microbial communities should be included in fate and effects assessment. Acknowledgments

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