Impact of inorganic nitrogen additions on microbes in biological soil crusts

Impact of inorganic nitrogen additions on microbes in biological soil crusts

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Soil Biology & Biochemistry xxx (2015) 1e11

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Soil Biology & Biochemistry journal homepage: www.elsevier.com/locate/soilbio

Q4 Q3

Impact of inorganic nitrogen additions on microbes in biological soil crusts Jin Wang a, *, 1, Jingting Bao a, b, c, 1, Jieqiong Su a, Xinrong Li a, Guoxiong Chen a, Xiaofei Ma a a

Laboratory of Plant Stress Ecophysiology and Biotechnology/Shapotou Desert Experiment and Research Station, Cold and Arid Regions Environmental and Engineering Research Institute, Chinese Academy of Sciences, Lanzhou 730000, China School of Life Science and Engineering, Lanzhou University of Technology, Lanzhou 730050, China c University of Chinese Academy of Sciences, Beijing 100049, China b

a r t i c l e i n f o

a b s t r a c t

Article history: Received 10 January 2015 Received in revised form 29 May 2015 Accepted 2 June 2015 Available online xxx

Many studies have shown that changes in nitrogen (N) availability affect the diversity and composition of soil microbial community in a variety of terrestrial systems, but less is known about the responses of microbes specific to biological soil crusts (BSCs) to increasing N additions. After seven years of field experiment, the bacterial diversity in lichen-dominated crusts decreased linearly with increasing inorganic N additions (ambient N deposition; low N addition, 3.5 g N m2 y1; medium N addition, 7.0 g N m2 y1; high N addition, 14.0 g N m2 y1), whereas the fungal diversity exhibited a distinctive pattern, with the low N-added crust containing a higher diversity than the other crusts. Pyrosequencing data revealed that the bacterial community shifted to more Cyanobacteria with modest N additions (low N and medium N) and to more Actinobacteria and Proteobacteria and much less Cyanobacteria with excess N addition (high N). Our results suggest that soil pH, together with soil organic carbon (C), structures the bacterial communities with N additions. Among the fungal communities, the relative abundance of Ascomycota increased with modest N but decreased with excess N. However, increasing N additions favored Basidiomycota, which may be ascribed to increases in substrate availability with low lignin and high cellulose contents under elevated N conditions. Bacteria/fungi ratios were higher in the N-added samples than in the control, suggesting that the bacterial biomass tends to dominate over that of fungi in lichen-dominated crusts after N additions, which is especially evident in the excess N condition. Because bacteria and fungi are important components and important decomposers in BSCs, the alterations of the bacterial and fungal communities may have implications in the formation and persistence of BSCs and the cycling and storage of C in desert ecosystems. © 2015 Published by Elsevier Ltd.

Keywords: Biological soil crusts Nitrogen addition Bacteria Fungi Pyrosequencing

1. Introduction Nitrogen (N) is a key element controlling the species composition, diversity and productivity of many terrestrial ecosystems (Zechmeister-Boltenstern et al., 2011). Over the past century, atmospheric deposition of reactive N (mainly nitrogen oxide and ammonia) has increased three- to fivefold (IPCC, 2007), and the atmospheric N deposition in terrestrial ecosystems is predicted to further increase by 250% over the next century (Lamarque et al.,

* Corresponding author. Tel.: þ86 9314967187. E-mail address: [email protected] (J. Wang). 1 J. Wang and J. Bao equally contributed to this work as first authors.

2005). It is well established that elevated N additions to ecosystems have wide-ranging consequences for the environment, including climate change, the emissions of greenhouse gases, species loss and even human health threats (Nemergut et al., 2008; Ramirez et al., 2010). The effects of elevated N deposition on altering the primary productivity in ecosystems have been extensively studied, and it is widely accepted that increases in N deposition can lead to increased C storage in the form of plant biomass and that higher N inputs can drive shifts in plant species composition, with growing evidence of a general trend towards a loss of diversity (Clark et al., 2007). Several studies have suggested that increased N may alter the microbial community structure and diversity and subsequent ecological function (Waldrop et al., 2004; Allison et al., 2008;

http://dx.doi.org/10.1016/j.soilbio.2015.06.004 0038-0717/© 2015 Published by Elsevier Ltd.

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Campbell et al., 2010). Amongst other changes, a shift towards bacteria-dominated microbial communities is expected (Tietema, 1998). Changes in the diversity and composition of soil microbial community after N addition have recently received increased attention in ecosystems, including forests (Entwistle et al., 2013), steppes (Zhang et al., 2008), grasslands and agriculture fields (Ramirez et al., 2010), and alpine tundras (Nemergut et al., 2008). A consensus of recent studies suggests that excess soil N reduces microbial biomass and activity and decreases the soil microbial respiration levels (Ramirez et al., 2010, 2012). To our knowledge, there are few consistent microbial responses to N addition across ecosystems. For example, experimental atmospheric N deposition seems to have no effect on the abundance of Actinobacteria on the forest floor (Eisenlord and Zak, 2010), while Ramirez et al. (2010) showed that Actinobacteria increased with N addition in both grasslands and agriculture fields. In addition, N manipulation variously reported increases, decreases or no net changes in the diversity and structure of fungal communities (Johnson, 1993; Frey et al., 2004; Jumpponen and Johnson, 2005; Allison et al., 2007). These changes may be ascribed to the different amounts and durations of N treatments, as meta-analyses showed that declines in the abundances of microbes (e.g., bacteria and fungi) and the changes in the microbial community structure were more evident with greater durations and amounts of N added (Treseder, 2008). Alternatively, each soil's unique set of starting characteristics (e.g., distinct edaphic characteristics and microbial communities) may induce very different, unquantifiable impacts of N enrichment on soil responses (Zeglin et al., 2007). Biological soil crusts (BSCs) are formed by communities of microorganisms that bind together the surface soil. Cyanobacteria, eukaryotic microalgae, fungi, mosses, bacteria and archaea are involved in the assembly of BSCs (Bates et al., 2010). BSCs are widespread in arid and semi-arid lands, and approximately onethird of the Earth's terrestrial surface is arid or semi-arid land where BSC coverage can be as high as 70% in some areas (Belnap and Eldridge, 2001). The ecological roles of BSCs (e.g., soil hydrological, soil biological and geochemical processes and ecological rehabilitation) have been well documented (Belnap, 2006; Bowker, 2007; Li et al., 2007; Moquin et al., 2012). Regarding the biological process of BSCs formation and persistence, it is crucial to keep a balance between microbial diversity, community structure and microbial abundance in the topsoil layer. If the soil microbes inside are disrupted in some way, it could have unexpected impacts on the ecological role and function of the BSCs. Therefore, our explanatory and predictive abilities with regard to the N management and preservation of BSCs must be based on a thorough mechanistic understanding of microbial responses to N constraints. However, few published studies (Porras-Alfaro et al., 2011) have analyzed BSC microbial community shifts with N addition. As bacteria and fungi are two of the most ecologically important components of BSCs (Gundlapally and Garcia-Pichel, 2006; Green et al., 2008) and most arid and semi-arid ecosystems are N-limited (McCalley and Sparks, 2009), there is good reason to suspect that increased N addition has the potential to alter the diversity and structure of the microbial community (e.g., shifts in the dominant phylotypes of bacteria and fungi) and/or alter the microbial abundance, especially the bacteria to fungi ratio. Either of these changes will very likely affect the microstructure and subsequent functional roles of the BSCs. To better understand how N additions impact the soil crust microbial diversity and community composition, we performed a manipulative field experiment with four N addition levels (0, 3.5, 7.0 and 14.0 g N m2 y1) on lichen-dominated crusts in the Tengger desert since 2007. Using N additions that include rates that few, if any, soils are likely to experience, we are able to determine

how the possible elimination of N limitation or excess N may alter microbial communities, the threshold of these responses, and possible mechanisms driving these responses (Ramirez et al., 2010). We measured the shifts of bacterial/fungal communities by 454 pyrosequencing and the changes of absolute bacterial/fungal abundance by quantitative real-time PCR (qPCR) and aimed to assess 1) whether the diversity or composition of bacterial and fungal communities in BSCs will show differential responses to N additions 2) whether the shifts of bacteria to fungi ratio are associated with the increasing of N additions and 3) the N addition thresholds of altering microbial communities. Understanding how N additions alter the composition of microbes specific to BSCs is critical to accurately predict terrestrial ecosystem responses and to identify approaches for ameliorating the negative effects of adverse environments. 2. Materials and methods 2.1. Study site and experimental design A simulated N deposition experiment was conducted starting in 2007 in a field with widespread lichen-dominated crusts as part of an existing experiment for assessing the long-term impacts of N additions on plant species diversity, productivity and dynamics (Su et al., 2013). This field is located in the Cuiliugou region (37 250 N, 104 350 E) on the southeastern fringe of Tengger desert in China) and has a crust coverage of greater than 90% (mainly Endocarpon pusillum Hedwig lichen-dominated crusts). The soils are wellbuffered alkaline sierozem and are thus only slightly prone to acidification. N in the form of NH4NO3 was added homogeneously to plots (1.0  1.0 m) covered with Endocarpon pusillum Hedwig lichen-dominated crusts without macroscopic mosses at rates of 0 (ambient N deposition, control), 3.5 (low N), 7.0 (medium N) and 14.0 g N m2 y1 (high N). Each plot was surrounded with an at least 1-m buffer zone that received the same N deposition. N was added in solution twice per year, half in mid-May and half in midJuly. Each treatment was repeated eight times (eight plots). To assure that N was the only limiting nutrient (Tilman, 1987), we added phosphorus (5 g P2O5 m2 y1) and some trace elements (Zn, Mn, B, Mo and Co) per year (Su et al., 2013), which are based on the soil census data (the PhD thesis of Jieqiong Su, unpublished data). All 32 plots were distributed randomly across an area of 20 m  50 m. 2.2. Soil sample collection and soil characterization In late October of 2013, four soil cores (5-cm depth and 3.5-cm diameter, with crust layers) from each of eight plots were sampled individually using a sterile trowel, and thus 32 soil cores were taken for each N treatment. All soil samples were randomly divided into duplicate aliquots in the field: one for nucleic acid-based molecular analysis and the other for soil physicochemical analysis. For the nucleic acid-based molecular analysis, the upper crust layers of the replicate plots were bulked and thoroughly mixed in the field to form a composite sample. Approximately 5 g of each composited sample was kept in a cooler box for transport to the laboratory and then stored at 70  C prior to DNA extraction (DNA was extracted within a month after sampling). For the soil physicochemical analysis, the total N was measured with a Kjeltec System 2300 €gana €s, Sweden); the soil available N was distilling unit (Tecator, Ho determined by the alkali diffusion method, the NO3  eN and NH4 þ eN contents were analyzed colorimetrically after the extraction of the fresh soil with 2 M KCl; available phosphorus (P) was extracted with 2 M ammonium acetate at pH 7.0; soil organic carbon (C) was measured by the Walkley-Black method; the soil pH

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was determined using a 1:5 (w/v) suspension of soil in distilled water and the soil water content in each plot was measured synchronously to soil sampling by the gravimetric method.

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were of the appropriate size. For each soil sample, the qPCR reactions were repeated four times. 2.5. Bioinformatics and statistical analysis

2.3. PCR and 454 pyrosequencing For this study, we focused on intersite variability, not the variability in microbial communities within individual plots, and thus, the DNA was extracted and amplified from the upper crust layers of a single, composite sample for each treatment. The four composites of 0, 3.5, 7.0 and 14.0 g N m2 y1 added soil crusts are referred to as NBC, NBM, NBL and NBH (for bacteria analysis) or NFC, NFM, NFL and NFH (for fungi analysis), respectively. Total DNA was isolated from 0.5 g of each soil sample using the PowerSoil DNA isolation kit (MOBIO Laboratories, Carlsbad, CA) according to the manufacturer's instructions. For each sample, DNA was extracted in triplicate to avoid bias, and the extracts from the same sample were pooled. The obtained DNA was quantified and examined for purity with NanoDrop 2000c (Thermo-Fisher Scientific). Amplification was performed in triplicate using a GeneAmp 9700 PCR system (Applied Biosystems) with TransStart Fastpfu DNA Polymerase (TransGen, China). Primers and barcodes are listed in Table S1. The 20-mL PCR reaction mixtures contained 5 FastPfu Buffer (4 mL), 2.5 mM dNTPs (2 mL), 5 mM aliquots of each primer (0.4 mL for bacteria/0.8 mL for fungi), FastPfu Polymerase (0.4 mL), 1 mL of template DNA (10 ng) and diethylpyrocarbonate-treated water. The reaction mixtures were amplified for 2 min at 95  C followed by 25 (for bacteria)/30 (for fungi) cycles at 95  C for 30 s, 55  C for 30 s, and 72  C for 30 s, with a final extension at 72  C for 5 min. The triplicate reaction products from each soil sample were pooled, and the pooled PCR products (amplicons) were purified with a DNA gel extraction kit (Axygen, China). The DNA concentration of each PCR product was determined using a QuantiFluor™ST dsDNA System (Promega) prior to sequencing. Following quantitation, the amplicons from each reaction mixture were pooled in equimolar ratios based on concentration and subjected to emulsion PCR to generate amplicon libraries, as recommended by 454 Life Sciences. Amplicon pyrosequencing was performed from the A-end using a 454/Roche A sequencing primer kit on a Roche Genome Sequencer GS FLX Titanium platform at Majorbio Bio-Pharm Technology Co., Ltd., Shanghai, China (http://www.majorbio.com). 2.4. Real-time PCR Quantitative real-time PCR (qPCR) was performed to determine the relative 16S rRNA and ITS rRNA gene abundance. We used the primer sets described by Fierer et al. (2005) for quantifying the total bacterial (Eub338/Eub518) and fungal (ITS1f/5.8S) populations (Table S1). The standard templates were made from 10-fold dilutions of linearized plasmids containing the gene fragment of interest that was cloned from amplified pure culture DNA (16S rRNA gene, five orders of magnitudes were 4.75  102e4.75  106 copies, Y ¼ 2.750  log (X) þ 33.91, R2 ¼ 0.987, efficiency ¼ 101.0%; ITS rRNA gene, five orders of magnitudes were 6.06  103e6.06  107 copies, Y ¼ 3.154  log (X) þ 34.77, R2 ¼ 0.991, efficiency ¼ 107.5%). The 20-mL PCR reaction mixtures contained 10 mL of Bestar™ SybrGreen qPCR Master Mix (DBI Bioscience), 0.5 mL each of 10 mmol/L forward and reverse primers, 1 mL of total DNA template (diluted to 10 ng/mL in advance), and 8 mL of sterile and DNA-free water. The reaction was conducted on a Stratagene Mx3000P Real-time PCR system (Stratagene, Agilent Technologies Inc., USA) using the following program: 95  C for 1 min followed by 40 cycles of 95  C for 5 s, 52  C for 15 s and 72  C for 15 s. The melting curve was obtained to confirm that the amplified products

Data preprocessing was performed mainly using the MOTHUR (version 1.25.1) program (Schloss et al., 2009). After sequencing, the original data were sorted into valid reads complying with the following rules: each pyrosequencing read containing a primer sequence should have no ambiguous bases and should match the primer and one of the barcode sequences used. To obtain reads of high quality for the down-stream analysis, we eliminated reads of less than 200 bp in length and with a quality score <25 using Seqclean (http://sourceforge.net/projects/seqclean/ & http://www. mothur.org/wiki/Main_Page). Then, the trimmed sequences were achieved after chimeras were removed using Chimera.uchime (MOTHUR). The bacterial sequences were aligned with the ribosomal sequence database in the SILVA library (SILVA 16S/18S, SSU. Version 111; http://www.arb-silva.de) using the kmer searching method (http://www.mothur.org/wiki/Align.seqs). The fungal sequences were aligned with all fungal sequences in the NCBI nt database, and the nonfungal-related reads were eliminated. Then, the fungal sequences were clustered to OTUs (operational taxonomic units) at 97% sequence identity using the MOTHUR program. The coverage percentage, community richness and diversity indices (Chao1 estimator, ACE and Shannon index), rarefaction curves, Metastats and Libshuff analyses were performed by the MOTHUR program, and the samples were clustered together using the unweighted pair group method with the arithmetic mean (UPGMA) algorithm by the MOTHUR program as well. One-way ANOVA and least significant difference (LSD) analysis were used to determine the effect of treatments on the crust microbial communities. All raw sequence reads are archived at the NCBI sequence Read Archive under accession numbers SRR1738247 (for bacteria), SRR1738248 and SRR1738249 (for fungi). 3. Results 3.1. Soil physicochemical characteristics The soil total N content as well as the total organic C content was significantly (P < 0.05 in all cases) higher in the N-added crusts than in the control crust (Table 1). Available N is defined as supplied N plus inorganic soil N. The available N and NO3  eN contents significantly increased in the medium-N and high-N crusts, with no significant difference (P > 0.05 in all cases) between the control and low-N crusts. The soil C/N ratios distinctly decreased from 8.62 to 6.49, while the NH4 þ eN content significantly increased from 0.58 to 4.66 mg/kg soil across the N additions. The soil pH decreased significantly across the N additions, but not dramatically, because of the well-buffered alkaline sierozem composition. The total P content and soil water content did not change significantly across the N additions. 3.2. Microbial a-diversity Four libraries of bacterial 16S rRNA genes and four libraries of eukaryotic ITS1F/ITS4 rRNA genes were constructed by 454 pyrosequencing. In total, 30,484 bacterial trimmed sequences and 22,714 ITS trimmed sequences (nonfungal reads removed) were retrieved with average lengths of 454 bp and 545 bp, respectively. Rarefaction analysis revealed that the control crust had the most bacterial OTUs, whereas the crust that received the high N contained the fewest bacterial OTUs. However, more fungal OTUs were observed with the high N addition compared to the other

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Table 1 Physico-chemical properties of seven years of increasing N-added soils (means ± s.d., n ¼ 5). The amounts of the added N were control (ambient N deposition), low (3.5 g N m2 y1), medium (7.0 g N m2 y1) and high (14.0 g N m2 y1), respectively. N-added Total N (g/Kg) Total P (g/Kg) Available N (mg/Kg) NO3  eN (mg/Kg) NH4 þ eN (mg/Kg) Organic C (g/Kg) C:N ratio Control Low Medium High

0.34 0.42 0.54 0.68

± ± ± ±

0.03 0.01 0.02 0.04

c b a a

0.34 0.39 0.41 0.43

± ± ± ±

0.02 0.01 0.02 0.01

a a a a

14.98 18.44 22.28 32.88

± ± ± ±

0.64 1.84 2.06 1.13

c c b a

2.47 2.66 3.86 7.14

± ± ± ±

0.21 0.22 0.23 0.66

c c b a

0.58 1.07 2.15 4.66

± ± ± ±

0.02 0.11 0.19 0.17

c bc b a

2.97 3.42 3.76 4.41

± ± ± ±

0.6 0.4 0.9 0.8

c b b a

8.62 8.14 7.01 6.49

± ± ± ±

0.4 0.5 0.8 0.6

pH a b c c

7.77 7.48 7.41 7.31

Water content (%) ± ± ± ±

0.04 0.03 0.05 0.02

a b b c

2.82 2.50 2.51 2.27

± ± ± ±

0.13 0.27 0.17 0.33

a a a a

Different letters indicate significant differences between treatments at P < 0.05.

treatments (Fig. 1). We calculated the diversity metrics for a randomly selected subset of 5500 sequences for bacteria and 3800 sequences for fungi per sample. For bacteria, the number of OTUs ranged from 2403 to 1915 across the N additions. The bacterial

community of the control was the most diverse, and the lowest OTU diversity was found in the sample that received the high N. Counter to our expectation, the fungal community of the high-N crust was the richest, with 498 OTUs. The fungal community of the control was the most diverse, while the lowest OTU diversity was present in the low-N crust. ACE and Chao1, two estimators of diversity, revealed that the control and low-N crusts contained higher bacterial and fungal diversities than the other crusts (Table 2). 3.3. Shifts of relative abundance of microbial taxa

Fig. 1. Rarefaction curves of the 16S rDNA libraries (A) and ITS rDNA libraries (B) based on 97% similarity. NBC/NFC, NBM/NFM, NBL/NFL and NBH/NFH mean bacterial/fungal samples received ambient N deposition, low N addition (3.5 g N m2 y1), medium N addition (7.0 g N m2 y1) and high N addition (14.0 g N m2 y1), respectively.

A total of 19 bacterial and 8 fungal phyla were retrieved at genetic distances of 3%. In the bacterial community, the highabundance phyla (>10% of total OTUs) included Actinobacteria, Chloroflexi, Cyanobacteria and Proteobacteria, and the lowabundance phyla (>1% of total OTUs) included Acidobacteria, Armatimonadetes, Bacteroidetes, Gemmatimonadetes and Planctomycetes. The high-abundance phyla and low-abundance phyla dominated the prokaryotic diversity. The other 10 phyla were all <1% of total OTUs, and as result, were removed from further analysis (Fig. 2A). The high-abundance phyla in the fungal community were only Ascomycota and no_rank_Fungi, whereas the lowabundance phyla consisted of Basidiomycota and unclassified_Fungi. The distribution of OTUs is illustrated in Fig. 2B. The other four phyla were all <1% of total OTUs and were not further analyzed. The bacterial community composition in the BSCs shifted in response to N additions (Fig. 3A). In the control crust (NBC), Proteobacteria was the most abundant phylum, accounting for 25.0% of the total reads, among which Alphaproteobacteria was prevalent, and relatively low abundances of Betaproteobacteria and Deltaproteobacteria (1.0e3.1%) were also presented, as well as a very small proportion of Gammaproteobacteria (approximately 0.2%). Cyanobacteria (19.5%) was the second most dominant phylum, followed by Actinobacteria (19.4%), Chloroflexi (12.2%), Bacteroidetes (6.6%), Acidobacteria (5.0%), Planctomycetes (4.6%), Armatimonadetes (3.1%), Gemmatimonadetes (3.0%) and other phyla. After seven years of N additions, Cyanobacteria became the most dominant phylum after the low N (NBM) and medium N (NBL) additions (27.7% and 33.4%, respectively) but dramatically decreased to 1.9% with the high N addition (NBH). The relative abundance of Actinobacteria, Chloroflexi and Alphaproteobacteria shared a similar trend to the N additions: all decreased slightly from the control to the medium N addition but sharply increased with the high N addition. Our results also showed that Acidobacteria and Planctomycetes continued to decline across the N additions, ranging from 5.0% to 2.0% and from 4.6% to 1.6%, respectively. The relative abundance of Bacteroidetes, Armatimonadetes and Gemmatimonadetes did not change much across the N additions. At the genus level, the numbers of detected bacterial genera were 275 in the NBC sample (control), 265 in the NBM sample (low N addition), 259 in the NBL sample (medium N addition) and 239 in the NBH sample (high N addition). The dominant genera

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Table 2 Diversity indices for the 16S rDNA sequences/ITS rDNA sequences after seven years of N additions. NBC/NFC, NBM/NFM, NBL/NFL and NBH/NFH mean bacterial/fungal samples received ambient N deposition, low N addition (3.5 g N m2 y1), medium N addition (7.0 g N m2 y1) and high N addition (14.0 g N m2 y1), respectively. Sample ID

NBC NBM NBL NBH NFC NFM NFL NFH

Trimmed reads

8119 8153 7122 7090 5464 6233 4324 6693

Normalized

5500 5500 5500 5500 3800 3800 3800 3800

OTU richness

OTU diversity

Good's coverage

Observed

ACE

Chao1

Shannon

Simpson

2403 2234 2150 1915 461 427 310 498

8118 7575 6320 5405 776 1126 715 860

5902 5464 4456 4075 788 869 572 847

7.26 7.02 6.85 6.84 4.28 3.75 3.81 4.26

0.0018 0.0032 0.0044 0.0043 0.0593 0.1354 0.0879 0.0473

0.7508 0.7692 0.7650 0.8126 0.9537 0.9574 0.9636 0.9584

Fig. 2. Abundant phyla (>10% of total OTUs) and low-abundance phyla (>1% of total OTUs) distributed in N-added samples for bacteria (A) and fungi (B), respectively. Data are defined at a 3% OTU genetic distance.

(abundance > 1% of total sequences) are shown in Fig. 3B. The most dominant genera were no_rank_Chloroflexi (9.8%), no_rank_Cyanobacteria (12.2%), Microcoleus (12.1%) and no_rank_Chloroflexi (12.4%) in samples NBC, NBM, NBL and NBH, respectively. Notably, the shifts in the Microcoleus and no_rank_Chloroflexi abundances (in the phyla Cyanobacteria and Chloroflexi) showed distinctly different response patterns between samples: Microcoleus increased from the control to medium N addition (6.1e12.1%) but decreased dramatically with the high N

addition (0.3%). However, no_rank_Chloroflexi decreased from the control to medium N addition (9.8e5.7%) but surprisingly increased to approximately 12.4% with the high N addition. Seven years of N additions also resulted in distinct shifts in the fungal communities specific to lichen-dominated crusts (Fig. 3C). Ascomycota predominated the fungal community in the control, accounting for 54.8% of the total reads, followed by a large proportion of nonassigned sequences (total 40.6%: no_rank_Fungi, 39.4%; unclassified_Fungi, 1.2%), and Basidiomycota (4.6%). The

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Fig. 3. Phylogenetic community composition. Relative abundance of dominant bacteria (>1% of total reads) (A) at the phylum level, with classes of Proteobacteria, (B) at the genus level, and dominant fungi (>1% of total reads) (C) at the phylum level, (D) at the genus level. NBC/NFC, NBM/NFM, NBL/NFL and NBH/NFH mean bacterial/fungal samples received ambient N deposition, low N addition (3.5 g N m2 y1), medium N addition (7.0 g N m2 y1) and high N addition (14.0 g N m2 y1), respectively.

phyla with the most prominent changes were Ascomycota and Basidiomycota. Ascomycota became the overwhelmingly dominant phyla with the low N (91.1%) and medium N (84.8%) additions but decreased to 21.9% with the high N addition, which was driven by a high percentage of OTUs appointed by a large proportion of nonassignable sequences (25.6% of total OTUs: no_rank_Fungi, 23.7%; unclassified_Fungi, 1.9%). The relative abundance of Basidiomycota distinctly increased from 4.6% to 7.3% across the N additions. Our results showed that Endocarpon (Ascomycota phylum) was the most dominant genus in the NFC sample (control), the NFM sample (low N addition) and the NFL sample (medium N addition), but in the NFH sample (high N addition) Fusarium (Ascomycota phylum) was the most dominant genus if non-assignable sequences were not counted. Among the three determined genera (all in the Ascomycota phylum), the abundance of Chalara decreased across the N additions, while Fusarium dramatically increased after the high N addition (NFH). Endocarpon increased from the control to the medium N addition (38.0e82.2%) but decreased to nearly 0 in NFH. The dominant genera (abundance >1% of total sequences) are shown in Fig. 3D. 3.4. Microbial b-diversity analysis Clustering of microbial communities in the crust samples by the UPGMA algorithm revealed that the bacterial community composition of NBC was much closer to that of NBM than to that of NBL or NBH, and the bacterial composition of NBH was quite different from those of other three samples. The same trend was observed in the fungal communities of NFC, NFM, NFL and NFH (Fig. S1). Furthermore, we assessed the differences in the microbial communities by the Libshuff analysis which compares the overall phylogenetic

diversity between microbial communities (Schloss, 2008). When every pair-wise comparison was made between all bacterial/fungal libraries (four for bacteria and the other four for fungi), all differences were significant (Table S2), indicating that these communities were significantly different. Metastats analysis was used for detecting differentially abundant features in samples (White et al., 2009) at the order level (for dominant groups, abundance >1% of total sequences) in 16S rRNA/ ITS rRNA gene libraries (Table S3). As shown in the bacterial samples, the Rhodobacterales abundance showed significant differences (P < 0.05 in all cases) between any two of the four samples except for NBC and NBL. No_rank_Actinobacteria were only sensitive to high N addition as no significant differences were found between NBC and NBM (P ¼ 0.764), NBC and NBL (P ¼ 0.176), or NBM and NBL (P ¼ 0.300). Notably, Oscillatoriales, as well as Prochlorales and Cyanobacteria_uncultured, showed significant differences (P < 0.05 in all cases) between any two of four samples, which indicates that these groups are very sensitive to N additions such that even a low N addition significantly changes their abundances. In the fungal samples, Cantharellales and Hypocreales had a similar response pattern to N additions: both were significantly different (P < 0.05 in all cases) between any two of the four samples except for NFC and NFL (P ¼ 1.000 and P ¼ 0.903). Pleosporales were only sensitive to the high N addition, as significant differences (P < 0.05 in all cases) were found between NFH and NFC, NFM, or NFL. Although Helotiales and Lichinales were sensitive to N additions, no significant differences were present between the medium and high N additions (P ¼ 0.166 and P ¼ 1.000, respectively). Additionally, the members of Verrucariales, no_rank_Fungi and unclassified_Fungi were very sensitive to N additions as each was significantly different (P < 0.05 in all cases) between any two of the four samples.

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et al., 2009; Seneviratne, 2009). In this work, the bacterial diversity decreased overall in N-added crusts, with the response linear to the N additions (Table 2 and Fig. 1A). In contrast to the prevalence of Cyanobacteria in cyanobacterial crusts (Nagy et al., 2005; Gundlapally and Garcia-Pichel, 2006; Abed et al., 2010), our observation that Proteobacteria (mainly Alphaproteobacteria) was the most abundant phylum in lichen-dominated crusts (in control) is consistent with previous lichen studies that used PCR-SSCP fingerprinting (Grube et al., 2009) and pyrosequencing (Maier et al., 2014). Cyanobacteria are major contributors to early-stage crust formation because they can stabilize soil, trap moisture, and import significant amounts of fixed carbon and nitrogen (Belnap, 2006; Housman et al., 2006). In the control crust, Cyanobacteria accounted for a large proportion of the bacterial sequences after Proteobacteria. The filamentous genus Microcoleus was highly abundant, which is consistent with the results in the Sonoran desert, Colorado Plateau and Gurbantunggut desert (Nagy et al., 2005; Gundlapally and Garcia-Pichel, 2006; Zhang et al., 2011). Meanwhile, a fairly high abundance of Actinobacteria (19.4%) was detected, which is similar to that found in cyanobacterial crusts and nonrhizobial soil communities (Nagy et al., 2005; Janssen, 2006), but distinct from bryophytic BSCs (Moquin et al., 2012). Unlike the studies of cyanobacterial crusts (Gundlapally and Garcia-Pichel, 2006), bryophytic crusts (Moquin et al., 2012) and lichendominated crusts (Psora decipiens-dominated crust and Toninia sedifolia-dominated crust, Maier et al., 2014), in which a relative low abundance of Chloroflexi was detected. However, in our crusts (Endocarpon pusillum-dominated crusts), this phylum contributed significantly to the bacterial community (12.2%), which is similar to the results of studies in the hyper-arid Atacama desert and mountain ‘barren’ soil (Freeman et al., 2009; Lacap et al., 2011). Members of Chloroflexi display a general adaptation to arid environments (Lacap et al., 2007, 2011), which seems to be important for the formation and persistence of BSCs in arid zones. Generally, soil pH strongly affects the microbial community structure (Lauber et al., 2009). Soil pH may be an important control on the shifts of Acidobacteria and Planctomycetes abundances, as both decreased corresponding with the changes of soil pH (Table 1 and Fig. 3A). The shifts of Acidobacteria can also be explained by the copiotrophic hypothesis (Fierer et al., 2007; Ramirez et al., 2010) in which the copiotrophic groups thrive in high C environments and oligotrophic groups survive best in low C environments. Acidobacteria and Bacteroidetes are often considered to be oligotrophic (Fierer et al., 2007; Davis et al., 2011). In the present study, the Acidobacteria abundance decreased as the total organic C content significantly increased with N additions (Table 1 and Fig. 3A). However, the results of the predicted oligotrophic group Bacteroidetes could not be adequately explained by this hypothesis as this phylum did not changed much across the N additions. Compared to other treatments, we observed distinct increases in the relative abundances of Betaproteobacteria, Actinobacteria and Chloroflexi with the high N addition. Betaproteobacteria and

3.5. Quantification of microbial abundance by qPCR The bacterial abundance in the control crust averaged approximately 1010 copies (16S rRNA gene) per gram of soil. The low N and medium N additions had no significant effect on the bacterial rRNA gene copy number (P ¼ 0.205 and P ¼ 0.356, respectively to control), while the high N addition significantly decreased the bacterial abundance (P ¼ 0.020, P ¼ 0.003 and P ¼ 0.005, respectively to the control, low N and medium N additions). The fungal ITS rDNA number in the control crust averaged approximately 108 copies per gram of soil. The low N and medium N additions significantly increased the abundance of fungi (P < 0.001, and P ¼ 0.003, respectively to control), while the high N addition greatly decreased the fungal abundance by an order of magnitude (P ¼ 0.038, P < 0.001, and P < 0.001, respectively to the control, low N and medium N additions). In all of the crust samples, we found that the bacterial abundance was much higher than that of fungi, and the high N addition significantly increased the bacteria/fungi ratio to approximately 200 (P ¼ 0.022, P ¼ 0.004 and P ¼ 0.005, respectively to the control, low N and medium N additions) (Table 3). 4. Discussion BSCs have been considered hotspots of biogeochemical inputs, as they mediate C and N cycling globally (Garcia-Pichel et al., 2003). Data from different BSCs indicate that crusts have up to 200% higher N content than uncrusted soils from the same site (Pointing and Belnap, 2012). However, nitrogen is still one of the limiting factors in BSCs due to increased microbial activity and the leaching of N into deeper soil layers (Schulz et al., 2013). According to the study of Lü and Tian (2007), the N deposition rate in the region of the Tengger desert was relatively low before 2007 (less than 1.0 g N m2 y1), but China's nitrogen deposition rate showed a higher mean value than those in the United States and Europe. The Tengger desert is very likely at the risk of suffering more serious N deposition in the future because of nearby intensive agriculture and recently developed industry. With the attempt to explore the threshold effect of added-N on microbes in BSCs, including under extreme conditions where the N deposition is very high, we used low (3.5 g N m2 y1), medium (7.0 g N m2 y1) and high (14.0 g N m2 y1) N addition rates. The present study examined how the elimination of N limitation and excess N addition altered microbial communities in lichen-dominated BSCs. The levels of N applied in this research are comparable to the maximum values observed in other dry regions (Bai et al., 2010; Harpole and Suding, 2011). 4.1. Changes of bacterial diversity and taxa across N additions Previous studies have shown that the overuse of inorganic nitrogen fertilizer can result in reduced microbial biodiversity (Hallin

Table 3 Absolute abundances of bacteria and fungi (copies of ribosomal genes per gram of soil) in lichen-dominated soil crusts quantified by qPCR and their paired ratios (means ± s.d., n ¼ 4). Lichen-dominated soil crusts have received seven years of N additions since 2007. The amounts of the added N were control (ambient N deposition), low (3.5 g N m2 y1), medium (7.0 g N m2 y1) and high (14.0 g N m2 y1). N-added Bacterial abundance

Control 1.43  1010e2.67 Low 1.88  1010e4.02 Medium 2.12  1010e3.36 High 2.23  109e4.99

Fungal abundance Average ± SD

Range    

1010 2.03  1010 1010 2.83  1010 1010 2.60  1010 109 3.36  109

± ± ± ±

Ratio (16S/ITS1f-5.8s) Average ± SD

Range 6.20 1.09 6.64 1.44

   

109 a 1010 a 109 a 109 b

1.85 6.52 5.57 1.28

   

108e3.36 108e9.86 108e8.24 107e2.03

   

108 108 108 107

2.53 8.01 6.64 1.64

   

108 108 108 107

± ± ± ±

7.68 1.70 1.41 3.76

Average ± SD

Range    

107 108 108 106

b a a c

4.27 1.91 2.57 1.10

   

101e1.12 101e5.25 101e5.50 102e3.10

   

102 101 101 102

8.73 3.71 4.08 2.15

   

101 101 101 102

± ± ± ±

3.88 1.69 1.46 1.00

   

101 101 101 102

b b b a

Different letters indicate significant differences between treatments at P < 0.05.

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Actinobacteria are usually predicted to be copiotrophic groups (Fierer et al., 2007), which increase in high C environments. Chloroflexi may also be copiotrophic as they are important organic matter decomposers and high C would spur their colonization (Freeman et al., 2009; Eisenlord and Zak, 2010). Surprisingly, our results showed that the shifts of Actinobacteria and Chloroflexi shared a similar trend with N additions, slightly decreasing from the control to the medium N addition but increasing dramatically with the high N addition, which suggests that soil pH and soil C (C mineralization) may control these microbial communities by stages. With the modest N addition (low N and medium N), the soil pH is the controlling factor, but with the high N addition, the soil C mineralization rate maybe the major control. Indeed, the soil C mineralization rate increased corresponding to the amount of N addition, without a significant difference between the control, low N and medium N additions but increasing significantly with the high N addition (Su et al., 2013 and our unpublished data). N saturation (excess N) may factor into the microbial responses, which are rarely linear (Ramirez et al., 2012), and the driving factors are therefore difficult to identify independently. Low doses of deposited N are important drivers for ecosystem development at the initial sites (Brankatschk et al., 2011). After seven years of low N or medium N application, the soil total N and available N increase to spur Cyanobacteria colonization and growth, and the ability of Cyanobacteria to trap nutrient-rich deposits via their exopolysaccharide sheath can facilitate this effect (Schulz et al., 2013). However, the high N addition may induce excess N for some groups, which is disadvantageous to Cyanobacteria (Fig. 3A). The shifts in the dominant genera across N additions were also pronounced. For example, among the dominant genera we detected, most members of Rubellimicrobium are positive for starch and the assimilation of cellulose (Dastager et al., 2008). The significant increase in this genus with the high N addition indicates that abundant N can stimulate the growth of cellulose-decomposing bacteria by relieving the N limitation for growth (Blackwood et al., 2007). Leptolyngbya and Microcoleus can produce extracellular exopolysaccharides (Pereira et al., 2009) which are important to the formation and persistence of BSCs. Relatively low doses of N addition (low N and medium N) promote the growth of both genera, but excess N (high N) evidently suppresses their colonization (Fig. 3B). The genera Rubrobacter and Microvirga display a general adaptation to thermal and arid environments (Ferreira et al., 1999; Laiz et al., 2009), and some species of Rubrobacter are known to be highly radiation-resistant (Yoshinaka et al., 1973). In addition, Actinoplanes is a mycelial genus (Vobis, 2006). The characteristics of the three above-mentioned genera are very important and helpful to the formation and persistence of BSCs in arid zones with enhanced ultraviolet radiation, although their abundances did not change much across N additions. 4.2. Shifts of fungal diversity and communities The internal transcribed spacer (ITS) region is considered the primary barcode marker for fungal identification (Schoch et al., 2012). Here, we used primers ITS1F/ITS4 to amplify the fungal ITS regions, which are widely used in sequencing-based fungal ecological studies (Yamamoto et al., 2012). Our results reveal that the fungal richness was higher than that obtained using the clone library and DGGE fingerprinting methods but similar to that obtained by pyrosequencing (Bent and Forney, 2008; Abed et al., 2013), which indicates that high-throughput 454-pyrosequencing can provide more information on the diversity of microorganisms than the DGGE and cloning approaches. The high-N added community, similar to the control community, was more diverse than

others. The ACE and Chao1 indices reveal that the high N-added crust contains a higher richness than do the other crusts (Table 2). These findings contrast with previous studies (Porras-Alfaro et al., 2011; Entwistle et al., 2013), which found no significant shifts in Chao1 richness and inverse Simpson diversity with N addition and significantly lower or no change in the Shannon diversity with N addition, but our work is partly consistent with the work by Robinson et al. (2004), in which low N addition (0.5 g N m2 y1) could marginally increase the fungal species richness in a high Arctic polar semidesert ecosystem. Interestingly, our results also revealed that a large proportion of the fungal diversity corresponds to nonassignable fungal sequences (no_rank_Fungi and unclassified_Fungi), which made up the second most common group next to Ascomycota in the control, low-N and medium-N crusts and mostly dominated in the high-N crusts. These numerous nonassignable sequences were also reported by Green et al. (2008), implying that many fungi specific to lichen-dominated BSCs are still unknown. When we performed the analysis of the diversity contributions of these sequences, no distinct differences were found from the control to the medium N addition. However, up to 70% of the sequences were non-assignable with the high N addition, which contributed 25.6% of the OTU diversity, suggesting that the increase in biodiversity in the high N-added crust was largely driven by the high proportion of nonassignable sequences. In agreement with a number of studies in deserts and grasslands (Green et al., 2008; Bates and Garcia-Pichel, 2009; Abed et al., 2013), our survey revealed that Ascomycota were prevalent in the fungal community in the control crust. The majority of fungal sequences belonged to the order Verrucariales, and approximately 38.0% of them were from the genus Endocarpon. This result is distinct from the studies by Porras-Alfaro et al. (2011) using the clone library method, Abed et al. (2013) using pyrosequencing and Bates et al. (2012) using DGGE fingerprinting, in which the order Pleosporales were widespread and frequently dominant, while this order exhibited a relatively low abundance in our work (1.4%). Nevertheless, it is not surprising that Endocarpon contribute the major fungal components in Endocarpon Pusillum-dominated crusts when we examined both the free-living fungi and lichenized fungi specific to our crusts, as Endocarpon Pusillum is a dominant lichen group that is widespread in the Tengger desert (Zhang and Wei, 2011) and our research region mainly comprises Endocarpon Pusillum-dominated crusts. Although no obvious regularity in the shifts of fungal diversity was found in response to the N additions, the four treatments resulted in four significantly different community structures (Table S2). Taxonomically, the relative abundance of Ascomycota increased with the low N and medium N additions, which is in agreement with previous studies (Allison et al., 2007, 2010; Nemergut et al., 2008), suggesting that the elimination of N limitation with the modest N addition is beneficial for the colonization of Ascomycota, whereas excess N was clearly harmful to the members of this phylum, especially to the genera of Chalara (Helotiales order) and Endocarpon (Verrucariales order) (Fig. 3D). In general, enhanced N availability should suppress lignin decay (Craine et al., 2007). Basidiomycota possess a suite of lignin oxidase enzymes, which are often considered the primary agents of lignin decomposition, and the decline of the relative abundance of these organisms is expected in N-added soils (Blackwood et al., 2007). However, our observed trends are in direct contrast with this prediction (Fig. 3C), suggesting that the shifts in the relative abundance of Basidiomycota were not associated with lignin decay. Indeed, crusted soils do not contain significant amounts of complex plantderived polymers (Bates and Garcia-Pichel, 2009). In our research region, the vegetation is dominated by sparsely distributed herbaceous species. Compared to the forest floor, the lignin content in

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crust soil is very much lower. It is known that the most effective litter decomposers with lignolytic capacities are found in the order Agaricales of the Basidiomycota phylum (Osono and Takeda, 2006;  Snajdr et al., 2010), but members of this order were found in very low abundances (all <1% of total sequences) across all of the N additions in the present study. Furthermore, the experimental N deposition did not consistently suppress the lignolytic Agaricales fungi (Entwistle et al., 2013). As lignin decay appeared to not affect the shifts of Basidiomycota, the relative abundance of this phylum may be influenced by the increase in crust soil organic matter induced by the added N, such as substrates with low lignin and high cellulose (Sinsabaugh et al., 2002), as enhanced N availability could stimulate the decomposition of low-lignin litter (Blackwood et al., 2007). 4.3. Abundance of bacteria and fungi The qPCR results revealed that the bacterial biomass was clearly dominant over fungi by a factor of approximately 10. The bacteria/ fungi ratio is one index that can improve our understanding of microbial community responses to N addition (Tietema, 1998; Frey et al., 2004). In this work, both the bacterial and fungal abundances increased after seven years of low N and medium N additions, but they decreased after receiving the high N addition. Notably, the increasing N additions trend toward more bacteria-dominated microbial communities, as revealed by higher bacteria/fungi ratio in the N-added crusts. Although there is some overlap in the physiological capabilities of bacteria and fungi, important differences are likely to affect their response to environmental changes. For example, the average C/N ratio in bacteria is approximately 4, and that in fungi is approximately 10 (De Deyn et al., 2008), suggesting that bacteria are able to use substrates of a lower C/N ratio than fungi. In this work, the C/N ratios in the upper soil layers decreased across the N additions, which would support more bacteria than fungi. Collectively, the shifts in the bacteria/fungi ratios suggest that the bacterial biomass tends to dominate over that of fungi in lichen-dominated BSCs after N additions, which is especially evident with excess N. 5. Conclusions Our results showed that the bacterial and fungal diversities and communities were shaped by seven years of increasing N additions. The bacterial diversity decreased with a response linear to the N additions, whereas the fungal diversity did not show an obvious pattern in response to the N additions. The high proportion of nonassignable sequences may be responsible for the increase in fungal diversity with the high N addition. The bacterial community shifted to more Cyanobacteria with the modest N additions, while more Actinobacteria and Proteobacteria and much less Cyanobacteria were present with excess N addition. Among the fungal communities, the abundance of Ascomycota increased with the modest N additions but decreased with the high N addition, while the Basidiomycota abundance increased after N additions. The absolute bacterial and fungal abundances increased with the modest N additions, whereas both were suppressed by excess N. In addition, a shift towards more bacteria-dominated microbial communities was supported by our qPCR data. Microbes have a central position in establishing crust communities (Garcia-Pichel and Wojciechowski, 2009). All of the above-mentioned alterations may have implications for the formation and persistence of BSCs and the cycling and storage of C in desert ecosystems. Still, these trends should be further verified by experiments in other ecosystems, such as in grassland and in cold or tropical deserts, to clarify whether there are consistent microbial responses to N additions

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66 67 68 69 70 71 Uncited reference 72 73 Gardes and Bruns, 1993, White et al., 1990, Wu et al., 2012. Q2 74 75 Acknowledgments 76 77 This work was financially supported by the State Key Develop78 ment Program for Basic Research of China (2013CB429906), the 79 National Natural Science Foundation of China (41201250), the 80 Natural Science Foundation of Gansu Province, China (1107RJYA068 81 and 145RJZA168), and the Innovation Research Group Fund of 82 Gansu Province, China (1308RJIA002). 83 84 Appendix A. Supplementary data 85 86 Supplementary data related to this article can be found at http:// 87 dx.doi.org/10.1016/j.soilbio.2015.06.004. 88 89 90 References 91 Abed, R.M.M., Al-Sadi, A.M., Al-Shehi, M., Al-Hinai, S., Robinson, M.D., 2013. Di92 versity of free-living and lichenized fungal communities in biological soil crusts 93 of the Sultanate of Oman and their role in improving soil properties. Soil 94 Biology and Biochemistry 57, 695e705. Abed, R.M.M., Kharusi, S.A., Schramm, A., Robinson, M.D., 2010. Bacterial diversity, 95 pigments and nitrogen fixation of biological desert crusts from the Sultanate of 96 Oman. FEMS Microbiology Ecology 72, 418e428. 97 Allison, S.D., Czimczik, C., Treseder, K., 2008. Microbial activity and soil respiration 98 under nitrogen addition in Alaskan boreal forest. Global Change Biology 14, 1156e1168. 99 Allison, S.D., Gartner, T.B., Mack, M.C., McGuire, K., Treseder, K., 2010. Nitrogen alters 100 carbon dynamics during early succession in boreal forest. Soil Biology and 101 Biochemistry 42, 1157e1164. Allison, S.D., Hanson, C.A., Treseder, K.K., 2007. Nitrogen fertilization reduces di102 versity and alters community structure of active fungi in boreal ecosystems. Soil 103 Biology and Biochemistry 39, 1878e1887. 104 Bai, Y.F., Wu, J.G., Clark, C.M., Naeem, S., Pan, Q.M., Huang, J.H., Zhang, L.X., Han, X.G., 2010. Tradeoffs and thresholds in the effects of nitrogen addition on biodiver105 sity and ecosystem functioning: evidence from inner Mongolia Grasslands. 106 Global Change Biology 16, 358e372. 107 Bates, S.T., Garcia-Pichel, F., 2009. A culture-independent study of free-living fungi in biological soil crusts of the Colorado Plateau: their diversity and relative 108 contribution to microbial biomass. Environmental Microbiology 11, 56e67. 109 Bates, S.T., Nash 3rd, T.H., Garcial-Pichel, F., 2012. Patterns of diversity for fungal 110 assemblages of biological soil crusts from the southwestern United States. Mycologia 104, 353e361. 111 Bates, S.T., Nash 3rd, T.H., Sweat, K.G., Garcial-Pichel, F., 2010. Fungal communities 112 of lichen-dominated biological soil crusts: diversity, relative microbial biomass, 113 and their relationship to disturbance and crust cover. Journal of Arid Environ114 ments 74, 1192e1199. Belnap, J., 2006. The potential roles of biological soil crusts in dryland hydrologic 115 cycles. Hydrological Processes 20, 3159e3178. 116 Belnap, J., Eldridge, D.J., 2001. Disturbance and recovery of biological soil crusts. In: 117 Belnap, J., Lange, O.L. (Eds.), Biological Soil Crusts: Structure, Function, and Management. Springer, Berlin, pp. 3e30. 118 Bent, S.J., Forney, L.J., 2008. The tragedy of the uncommon: understanding limita119 tions in the analysis of microbial diversity. The ISME Journal 2, 689e695. 120 Bowker, M.A., 2007. Biological soil crust rehabilitation in theory and practice: an underexploited opportunity. Restoration Ecology 15, 13e23. 121 €we, S., Kleineidam, K., Schloter, M., Zeyer, J., 2011. Abundances Brankatschk, R., To 122 and potential activities of nitrogen cycling microbial communities along a 123 chronosequence of a glacier forefield. The ISME Journal 5, 1025e1037. Campbell, B.J., Polson, S.W., Hanson, T.E., Mack, M.C., Schuur, E.A.G., 2010. The effect 124 of nutrient deposition on bacterial communities in Arctic tundra soil. Envi125 ronmental Microbiology 12, 1842e1854. 126 Clark, C.M., Cleland, E.E., Collins, S.L., Fargione, J.E., Gough, L., Gross, K.L., Pennings, S.C., Suding, K.N., Grace, J.B., 2007. Environmental and plant com127 munity determinants of species loss following nitrogen enrichment. Ecology 128 Letters 10, 596e607. 129 Craine, J.M., Morrow, C., Fierer, N., 2007. Microbial nitrogen limitation increases 130 decomposition. Ecology 88, 2105e2113. across ecosystems. Moreover, control experiments with standardized, laboratory conditions should be performed to eliminate other factors (e.g. indirect effects of N additions on plant C inputs) and focus on microbial responses only to N additions.

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