Effects of ammonium on the activity and community of methanotrophs in landfill biocover soils

Systematic and Applied Microbiology 37 (2014) 296–304

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Effects of ammonium on the activity and community of methanotrophs in landfill biocover soils Xuan Zhang, Jiao-Yan Kong, Fang-Fang Xia, Yao Su, Ruo He ∗ Department of Environmental Engineering, Zhejiang University, Hangzhou 310058, China

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Article history: Received 3 December 2013 Received in revised form 23 March 2014 Accepted 24 March 2014 Keywords: Methanotroph Waste biocover soil NH4 + -N addition Methane oxidation

a b s t r a c t The influence of NH4 + on microbial CH4 oxidation is still poorly understood in landfill cover soils. In this study, effects of NH4 + addition on the activity and community structure of methanotrophs were investigated in waste biocover soil (WBS) treated by a series of NH4 + -N contents (0, 100, 300, 600 and 1200 mg kg−1 ). The results showed that the addition of NH4 + -N ranging from 100 to 300 mg kg−1 could stimulate CH4 oxidation in the WBS samples at the first stage of activity, while the addition of an NH4 + N content of 600 mg kg−1 had an inhibitory effect on CH4 oxidation in the first 4 days. The decrease of CH4 oxidation rate observed in the last stage of activity could be caused by nitrogen limitation and/or exopolymeric substance accumulation. Type I methanotrophs Methylocaldum and Methylobacter, and type II methanotrophs (Methylocystis and Methylosinus) were abundant in the WBS samples. Of these, Methylocaldum was the main methanotroph in the original WBS. With incubation, a higher abundance of Methylobacter was observed in the treatments with NH4 + -N contents greater than 300 mg kg−1 , which suggested that NH4 + -N addition might lead to the dominance of Methylobacter in the WBS samples. Compared to type I methanotrophs, the abundance of type II methanotrophs Methylocystis and/or Methylosinus was lower in the original WBS sample. An increase in the abundance of Methylocystis and/or Methylosinus occurred in the last stage of activity, and was likely due to a nitrogen limitation condition. Redundancy analysis showed that NH4 + -N and the C/N ratio had a significant influence on the methanotrophic community in the WBS sample. © 2014 Elsevier GmbH. All rights reserved.

Introduction Aerobic CH4 oxidation in oxic layers of landfill cover soils plays a significant role in reducing CH4 emissions from landfills. CH4 oxidation in these soils is affected by environmental factors such as soil texture, pH, soil moisture content, CH4 and O2 supply, nutrients and temperature [34]. Nitrogen is an important nutrient for microorganisms that can affect methanotrophic activities, and it subsequently interferes with the capacity of CH4 oxidation in soil. Some studies have shown that higher NH4 + contents in soils tend to inhibit CH4 oxidation, due to substrate competition between NH3 and CH4 for the active site of methane monooxygenase (MMO), which catalyzes the oxidation of CH4 to methanol, and the toxicity of hydroxylamine and nitrite generated from NH3 oxidation [23,24,37–39]. Different amounts of NH4 + might contradict the effects of CH4 oxidation. The CH4 oxidation rate in landfill cover soils was shown to decrease linearly with the initial

∗ Corresponding author. Tel.: +86 571 88982221; fax: +86 571 88982221. E-mail address: [email protected] (R. He). http://dx.doi.org/10.1016/j.syapm.2014.03.003 0723-2020/© 2014 Elsevier GmbH. All rights reserved.

NH4 + content of the soil when the NH4 + -N content reached 25 mgN kg−1 [4]. Similar results were obtained by Scheutz and Kjeldsen [33] who showed that CH4 oxidation rates were unaltered when NH4 + -N amended soil was 14 mg-N kg−1 or below, whereas the oxidation rates decreased at a higher NH4 + -N content. Some studies, on the other hand, indicated that NH4 + -N application could stimulate growth and activity of methanotrophs in landfill cover soils and rice fields [3,10,26,31]. When CH4 flux is high enough to support the growth of methanotrophs, NH4 + can act more as a nutrient than as an inhibitor and strengthen the biological CH4 sink [37]. The stimulative effect of NH4 + on CH4 oxidation is also explained by the competition for nitrogen between the plant and the rhizosphere microbial community, thereby reducing the effective NH4 + concentration below a threshold that is inhibitory to methanotrophic activity [2]. In addition, since aerobic methanotrophs require oxygen for growth, sufficient oxygen generated from high plant density with nitrogen addition can stimulate CH4 oxidation [2]. Aerobic methanotrophs are the major mediator in mitigating CH4 emission from landfills, and most of them belong to Proteobacteria that can be classified into two major groups, type I and type

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II, based on their cell morphology, ultrastructure, phylogeny and metabolic pathways [14,36]. Nitrogen content has an important effect on the distribution of methanotrophs in the natural environment [34]. The activity and community of methanotrophs in a landfill cover soil with NH4 + addition has been hypothesized to occur in three stages, according to the growth and activity of methanotrophs with relatively high CH4 mixing ratios (>1%) [9]. In the first stage, methanotrophs have a rapid growth rate, with higher rates probably obtained for type I methanotrophs. The second stage represents a decline in the methanotrophic activity, which might be caused by nitrogen limitation conditions for type I methanotrophs. In the third stage, a new growth phase can be observed that might be dominated by N2 -fixing type II methanotrophs. Compared to type I methanotrophs, type II methanotrophs can survive in a nitrogen-limited environment [12,22], which could be due to their nitrogen fixation capacity. Biocover soils, such as compost, waste biocover soil (WBS) and mineralized waste, have high organic matter content, welldistributed particle size and active microorganism activity and have been demonstrated to be good alternative covers for mitigating CH4 emission from landfills [5,17,33]. Ammonium is an important compound generated from the decomposition of deposited waste in landfills, especially in bioreactor landfills with leachate recycling [7,16]. The immediate effect of nitrogen (NH4 + -N and NO3 − -N) addition on CH4 oxidation and N2 O emission in biocover soils has been investigated for short periods (within 150 h) in batch experiments [42,44]. However, there is little information on the responses of methanotrophic communities and their activity to NH4 + -N application in landfill biocover soils. Therefore, the objective of this study was to test the effects of NH4 + -N addition on the activity of the methanotroph community in landfill biocover soils. Batch experiments were conducted with a series of NH4 + -N contents (0, 100, 300, 600 and 1200 mg kg−1 ) in WBS. NH4 + -N, NO3 − -N, total nitrogen (TN), total organic carbon (TOC) contents and the C/N ratio were determined during incubation. Quantitative PCR (Q-PCR), terminal restriction fragment length polymorphism (T-RFLP) and cloning of pmoA (encoding a subunit of the particulate MMO) were applied to analyze the identity and diversity of methanotrophs and their response to NH4 + -N addition.

Materials and methods Landfill cover soil microcosm The WBS used in this study was taken from an organic waste landfill bioreactor (2 m3 ) in a village located in Xindeng town, Zhejiang Province. After removing large particles, the soils were air-dried and sieved through a 4 mm mesh. The particle composition of the experimental WBS samples was 61% of 2–4 mm, 33% of 0.02–2 mm, 5% of 0.002–0.02 mm and 1% of <0.002 mm. The pH value was 7.7. The TN, NO3 − -N, NO2 − -N and NH4 + -N contents of the WBS samples were 710.0 mg kg−1 , 674.3 mg kg−1 , 0.9 mg kg−1 and 33.4 mg kg−1 , respectively. Approximately 70 g of the air-dried experimental WBS were placed into 400 mL serum bottles. The WBS was adjusted to a water content of 45% (w/w), at which the WBS was reported to have the highest CH4 oxidation activity [42]. The NH4 Cl solution was added to the WBS samples by adding quantities of 100 mg-N kg−1 , 300 mg-N kg−1 , 600 mg-N kg−1 and 1200 mg-N kg−1 (denoted as WN100, WN300, WN600 and WN1200, respectively). The treatment without NH4 Cl addition was denoted as CK. The treatments with sterilized soils and NaN3 (0.13 mg g−1 dry weight) were used as non-microbial controls to check if CH4 disappearance occurred. All tests were performed in triplicate.

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After the adjustment of the soil water and NH4 + -N contents, the serum bottles were sealed with butyl rubber stoppers, and they were then covered with cling film and allowed to equilibrate overnight (∼approximately 12 h) at 30 ◦ C. A corresponding volume of air was taken from the serum bottles prior to injecting simulated landfill gas (LFG). Simulated LFG (CH4 :CO2 = 1:1; v/v) was injected into the serum bottles at CH4 and CO2 concentrations of 10% (v/v). High purity O2 was injected into the serum bottles at a concentration of ∼21% (v/v). Gas samples were withdrawn periodically from the headspace of the serum bottles to measure CH4 , CO2 and O2 concentrations. The serum bottles were flushed with air and the initial concentrations of CH4 , CO2 and O2 were resupplied every 12 h to reestablish their initial concentrations. The whole experiment lasted for 56 days. Soil sampling and analyses Soil samples were withdrawn from the serum bottles after 7, 15, 30 and 56 days incubation. They were immediately used to measure soil water moisture, the contents of TN, NO3 − -N, NH4 + -N, TOC and extracellular polymeric substances (EPS), including extracellular protein (ECP) and extracellular polysaccharide (ECPS), and the remaining samples were stored at −20 ◦ C for subsequent molecular analysis. The subsample withdrawn at the end of the experiment was used to monitor soil pH value. The TN, NH4 + -N, NO3 − -N and TOC contents were determined using the methods described by Bao [1]. The soil water content was measured by the loss of soil weight after drying to a constant weight at 105 ◦ C. EPS was measured using the method modified from McSwain et al. [29]. Approximately 2.0 g of fresh soil were placed into 50 mL centrifuge tubes containing 20 mL sterile water. After being centrifuged at 4 ◦ C and 4000 rpm for 5 min, the supernatant was discarded and the clean soil sample was used to extract EPS using 15 mL of 1 M NaOH solution heated in a water bath at 80 ◦ C for 30 min. The extract was recovered by centrifuging at 10,000 rpm for 10 min, and then the supernatant was used to detect protein and polysaccharide by using the Bradford [6] and phenol-sulfate methods [11], respectively. CH4 oxidation rate measurement CH4 oxidation rate was measured using the method described by Wang et al. [42]. In brief, after CH4 injection, four gas samples were taken periodically from the headspace of the serum bottles. Then, the CH4 oxidation rates were calculated by the regression of the amount of CH4 loss (i.e. the amount of CH4 loss in a non-microbial control was subtracted from that in the treatment) against time. The CH4 concentrations were measured using a gas chromatograph equipped with a thermal conductivity detector (TCD). The gas chromatography conditions for the TCD were as follows: two 2 m stainless steel columns filled with GDX-103 (80/100 mesh) and 5A molecular sieves, respectively; the temperatures of the oven, injector and detector were 90 ◦ C, 100 ◦ C and 100 ◦ C, respectively. Hydrogen with a 99.999% purity was used as the carrier gas and it was introduced at a flow rate of 20 mL min−1 . DNA extraction, Q-PCR and T-RFLP analysis DNA was extracted from 0.5 g soil samples with the E.Z.N.A.TM Soil DNA Kit (Omega Bio-Tek, Inc., Norcross, USA). The experimental triplicates of each soil sample were used for DNA extraction, respectively. DNA was quantified using the Nanodrop ND-1000 spectrophotometer, and it was then mixed in equal amounts to obtain the DNA for each treatment and sampling time point sample for Q-PCR and T-RFLP analysis. The pmoA gene was amplified using

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the primers A189F [21] and mb661R [8]. Q-PCR and PCR for T-RFLP analysis were performed according to the method described by Kong et al. [25]. PCR products for T-RFLP analysis were purified using the AxyPrepTMDNA Gel Extraction Kit (Axygen Scientific Inc. Union City, CA, USA) and quantified using the Nanodrop ND-1000 spectrophotometer. Then, 40 ng of purified PCR products were digested with restriction endonuclease MspI (Fermentas International Inc., ON, Burlington, Canada) for 3 h at 37 ◦ C and precipitated according to the procedure described by He et al. [18]. Pellets were resuspended in 1.0 ␮L H2 O, 9.5 ␮L deionized formamide and 0.5 ␮L of GeneScanTM 500LIZTM Size Standard (Applied Biosystems, Foster City, CA, USA), and then run on an ABI 3730XL Genetic Analyzer (Applied Biosystems, Foster City, CA, USA) at Sangon Biotech Co. Ltd. (Shanghai, China). T-RFLPs were analyzed as described by Kong et al. [25].

Cloning and phylogenetic analyses DNA extracted from all the soil samples was mixed in equal amounts to obtain the total DNA for all the soil samples, and DNA for the CK, WN100, WN300, WN600 and WN1200 soil samples on day 56, respectively, for PCR amplification. The PCR reaction mixture was identical to that of the T-RFLP analysis except that primer A189F was used instead of the 5 6-carboxyfluoresceinlabeled primer A189F. The thermal cycler conditions, PCR product purification and cloning were performed according to the method described by Kong et al. [25]. Clones of the total DNA were randomly selected for screening plasmid inserts and then 125 positive clones were sequenced at the Beijing Liuhehuada Gene Incorporation (China). DNA clones of the CK, WN100, WN300, WN600 and WN1200 soil samples on day 56, respectively, were obtained as described previously [25]. Positive clones were digested with restriction endonucleases HaeIII (Fermentas International Inc., Burlington, ON, Canada) and MspI. Each clone was assigned to an operational taxonomic unit (OTU) that represented a unique restriction fragment length polymorphism. From each OTU, 10% of the clones were sequenced and analyzed. To check the difference of T-RF sizes between the determined peak and the predicted data generated by in silico digestion of the cloning sequences, clones were chosen for conducting T-RFLP analysis. Raw pmoA sequences were first trimmed and primer sequences were removed using DNASTAR software (DNASTAR Inc., Madison, USA), and then were inspected for chimeras by searching for large sequence regions of unexpected nucleotide changes when compared with the reference sequences. The remaining sequences in each clone library were assembled with DNASTAR software with 97% sequence identity cut-off. In the clone library of the total DNA, one representative of each cluster was aligned with related sequences extracted from GenBank and translated to obtain deduced amino acid sequences. A phylogenetic tree was constructed using MEGA 5 software with the maximum-likelihood method [18]. Sequences in this study have been submitted to the GenBank database under accession numbers KF727479–KF727507 and KJ466865–KJ466895.

Data analysis Redundancy analysis (RDA) was carried out with the package CANOCO 4.5. The statistical significance of the relationship between the environmental variables and the variation in pmoA T-RFLP profiles was performed using the Monte Carlo test based on 499 random permutations.

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Fig. 1. Variation of CH4 oxidation rates over time in the WBS samples with NH4 + -N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 .

Results Effects of NH4 + -N addition on CH4 oxidation activity The CH4 oxidation rates in the five experimental treatments showed a similar trend that increased in the first stage at the first stage, reached the maximum values of 110–178 mg kg−1 h−1 between days 18 and 22, and then declined to lower values with a slight fluctuation until the end of the experiment (Fig. 1). Among the five treatments, the highest CH4 oxidation rates occurred in the WN100 treatment in the first 4 days. On day 4, the second-highest CH4 oxidation rate was observed in the CK and WN300 treatments with 32–35 mg kg−1 h−1 , followed by the WN600 treatment, and then the lowest rate was observed in the WN1200 treatment. After day 4, the CH4 oxidation rate increased rapidly in the WN300 treatment, and reached the peak value (135 mg kg−1 h−1 ) on day 11, after which it had a decreasing trend between days 11 and 25. The oxidation rate in the WN600 treatment also increased rapidly, and reached a high value of 143–157 mg kg−1 h−1 between days 15 and 22, but then decreased sharply to 31 mg kg−1 h−1 on day 29. Compared to the WN300 and WN600 treatments, the CH4 oxidation rates in the CK, WN100 and WN1200 treatments showed a slower increase and reached the maximum values on day 18 or 22. The variation of the CH4 oxidation rate in the WN1200 treatment was less compared to the others, and it had the lowest peak value of 110 mg kg−1 h−1 during the experiment. Between days 29 and 45, the CH4 oxidation rate in the WN1200 treatment was markedly higher than the others (P = 0–0.035). However, after day 45, there was no obvious difference in the CH4 oxidation rate between the five treatments (P = 0.089–0.914). Variation of NH4 + -N, NO3 − -N contents and C/N ratio The NH4 + -N and NO3 − -N contents of the original WBS were 33 mg kg−1 and 673 mg kg−1 , respectively (Fig. 2). With incubation, the NH4 + -N contents of WBS declined smoothly and almost disappeared in the CK treatment on day 30. The NO3 − -N contents of WBS also had a decreasing trend with increasing incubation time. Compared to the WN600 and WN1200 treatments, a more rapid decrease of NO3 − -N contents occurred in the CK, WN100 and WN300 treatments. On day 30, the NO3 − -N contents of WBS in these treatments were between 4.9 and 22.8 mg kg−1 , while they were greater than 102 mg kg−1 in the WN600 and WN1200 treatments. At the end of the experiment (day 56), the NH4 + -N and NO3 − -N contents in all experimental treatments were close to 0.

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Fig. 2. Variation of NH4 + -N, NO3 − -N and total nitrogen (TN), total organic carbon content (TOC) and C/N ratio over time in the WBS samples with NH4 + -N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 .

The TN content in the experimental treatments showed a similar variation trend to that of the NH4 + -N content. The TN content of the original WBS was 1284 mg kg−1 . With incubation, the TN contents in the treatments showed a rapid decrease in the first 30 days, especially in the treatments with NH4 + -N addition. After that, the TN contents decreased slowly and maintained similar values in all the experimental treatments on day 56. The TOC contents in the experimental treatments reduced rapidly from 56.9 to 33.1–40.2 g kg−1 in the first 7 days. After that, the TOC contents in each treatment decreased gradually with incubation. Statistical analyses showed that there was no significant difference in the TOC contents during incubation between the CK and the WN100 treatments (P ≥ 0.371). Compared to the CK treatment, the TOC content was lower in the WN300, WN600 and WN1200 treatments at the end of the experiment. The C/N ratio of the original WBS was 44. The C/N ratios of the WBS samples in the WN100, WN300, WN600 and WN1200 treatments after the addition of NH4 + -N were 41, 36, 30 and 23, respectively. With incubation, the C/N ratio had a generally increasing trend. However, the C/N ratio occasionally decreased due to a relatively larger decrease in TOC than in TN. The C/N ratio increased quickly and reached 119 and 95 in the CK and WN100 treatments, respectively, on day 30. At the end of the experiment, the C/N ratios were 130, 98, 85 and 88 in the WN100, WN300, WN600 and WN1200 treatments, respectively.

Variation of EPS content The ECPS and ECP contents of the original WBS were 379 mg kg−1 and 45 mg kg−1 , respectively (Fig. 3). With incubation,

the ECPS contents showed an increasing trend in the first 30 days. On day 7, the highest ECPS contents were observed in the WN100 treatment, followed by those in the WN600, WN300 and CK treatments, whereas the lowest occurred in the WN1200 treatment. NH4 + -N addition enhanced the secretion of ECPS. The ECPS contents reached the maximum values of 762 mg kg−1 , 803 mg kg−1 , 918 mg kg−1 , 978 mg kg−1 and 1526 mg kg−1 on day 30 in the CK, WN100, WN300, WN600 and WN1200 treatments, respectively. At the end of the experiment, the ECPS contents decreased slightly in all the experimental treatments. The ECP contents increased with time in all the experimental treatments. Compared to the treatments with NH4 + -N addition, the ECP content was lower in the CK during incubation. Among the four treatments with NH4 + -N addition, the highest ECP contents occurred in the WN1200 treatment, followed by the WN100, WN600 and WN300 treatments. At the end of the experiment, the ECP contents reached 559 mg kg−1 in the WN1200 treatment, which was almost twice as high as those in the CK, WN300 and WN600 treatments.

Overall methanotrophic community structure To assess the overall community structure of methanotrophs, DNA extracted from all the soil samples for the T-RFLP analysis was combined together and used to create a pmoA clone library. A total of 119 clones were obtained and the sequences were subjected to BLASTn searches using GenBank. Among them, 111 clones (93.3%) belonged to Methylocaldum-like species, of which 86 clones showed 93–99% sequence similarity to Methylocaldum tepidum (U89304); 13 clones showed 94–99% sequence similarity to Methylocaldum sp.

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Fig. 3. Extracellular polysaccharide (ECPS) and extracellular protein (ECP) contents over time in the WBS samples with NH4 + -N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 .

Fig. 4. Maximum-likelihood tree of deduced pmoA sequences amplified from the composite DNA of each individual sample. The scale bar represents 0.1 substitutions per nucleotide position. Bootstrap values greater than 50 are shown. The number of clones in the same operational taxonomic unit (OTU) analyzed by restriction fragment length polymorphism (RFLP) is shown in parentheses.

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Fig. 5. Quantitative real-time PCR (Q-PCR) of pmoA in the WBS samples with NH4 + N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 . Error bars denote the standard deviation of three replicates for the composite DNA of each treatment and sampling time point.

E10 A (AJ544091); 12 clones showed 90–99% sequence similarity to Methylocaldum sp. 05J-I-7 (EU275141) (Fig. 4). Five of the 119 clones (4.2%) showed 92–99% sequence similarity to Methylobacter sp. BB5.1 (AF016982) and Methylobacter sp. 5FB (AJ868410); two clones showed 91% sequence similarity to Methylococcaceae bacterium OS501 (AB636304); one clone showed 99% sequence similarity to type II methanotroph Methylocystis sp. 42/22 (AJ459009). Effect of NH4 + -N addition on diversity and abundance of methanotrophs Q-PCR analysis of pmoA showed that the abundance of methanotrophs was 1.03 × 106 copies g−1 in the original WBS (Fig. 5). With incubation, the abundance of methanotrophs in the experimental treatments increased to 2.38 × 107 –6.75 × 107 copies g−1 except for the WN1200 treatment that had an abundance of 1.1 × 106 copies g−1 on day 7. The highest abundance of methanotrophs in the WN100 treatment was observed on day 15 with an abundance of 4.25 × 108 copies g−1 , while the highest in the CK treatment occurred on day 30 with 4.18 × 108 copies g−1 . The abundance of methanotrophs in the WN300, WN600 and WN1200 treatments increased with time, except for a slight decrease in the WN300 and WN600 treatments on days 15 and 30, respectively. At the end of the experiment, the abundance of methanotrophs in the WN300, WN600 and WN1200 treatments reached maximums of 1.87 × 108 copies g−1 , 1.51 × 108 copies g−1 and 3.70 × 108 copies g−1 , respectively. The pmoA clone libraries for the DNA of the CK, WN100, WN300, WN600 and WN1200 soil samples on day 56 showed that Methylocaldum dominated in each library, and accounted for 74–93% (Fig. 6). Methylosinus was abundant in the CK treatment and reached 25.6% of the cloning sequences. With the increase of NH4 + -N content, the abundance of Methylosinus decreased, and was 0 in the WN600 and WN1200 treatments on day 56. Methylococcaceae was detected at the low abundance of 2.3–6.2% in the WN100 and WN300 treatments. Methylobacter was not detected in the CK and WN100 treatments on day 56, but it was present in the WN300, WN600 and WN1200 treatments. The highest abundance of Methylobacter (25.6%) was observed in the WN1200 treatments at the end of the experiment. Predictions of T-RF sizes generated by in silico digestion of the cloning sequence data showed that the 79-bp T-RF was associated with Methylocaldum and/or Methylococcaceae, the 244-bp T-RF was associated with Methylocystis and/or Methylosinus, and the 508 bp

Fig. 6. Relative abundance of pmoA cloned sequences from DNA of the WBS samples with NH4 + -N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 on day 56.

T-RF was related to Methylobacter (Figs. 4 and 6). Type I, including Methylocaldum, Methylococcaceae and Methylobacter, and type II methanotrophs (Methylocystis and Methylosinus) were abundant in the WBS samples, accounting for 87–99% of the total T-RF abundance, except in the WN300 treatment on day 30 that had an abundance of 70% (Fig. 7). In the original WBS sample, Methylocaldum and/or Methylococcaceae were the main methanotrophs, accounting for 88% of the total T-RF abundance. With incubation, the abundance of Methylocaldum and/or Methylococcaceae had a decreasing trend in all the experimental treatments on day 7. After that, the abundance of Methylocaldum and/or Methylococcaceae in the WN100 treatment remained relatively stable with a slightly fluctuating abundance ranging from 68% to 75%. The abundance of Methylocaldum and/or Methylococcaceae in the CK treatment increased after day 7, while it showed an increase in the WN300 and WN600 treatments after day 15. The abundance of Methylocaldum and/or Methylococcaceae in the WN1200 treatment continuously decreased to 22% until day 30, and then increased to 45% on day 56. The abundance of Methylobacter in the original WBS was 1.8%, and with incubation the abundance showed an increasing trend in all the experimental treatments. The abundance of Methylobacter in the CK treatment reached a maximum on day 7 with an abundance of 37%, while it reached the peak value in the WN100, WN300 and WN600 treatments on day 15 with abundances of 24%, 73% and 66%, respectively. The abundances of Methylobacter in the CK, WN100, WN300 and WN600 treatments decreased rapidly and had similar values of 7–11% at the end of the experiment. The abundance of Methylobacter in the WN1200 treatment showed an increasing trend, and reached the maximum value of 76% on day 30. Compared to type I methanotrophs, type II methanotrophs Methylocystis and/or Methylosinus were lower in the original WBS sample and the experimental WBS samples during the first 30 days with abundances of 0.2–1%, except for an abundance of 5% in the WN100 treatment on day 30. After day 30, the abundance of Methylocystis and/or Methylosinus showed an increasing trend, and reached 3–17% in the CK, WN100, WN300 and WN600 treatments, while it was low (almost 0) in the WN1200 treatment during the whole experiment.

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Fig. 8. Redundancy analysis ordination biplot of terminal restriction fragment length polymorphism (T-RFLP) profiles of pmoA amplicons digested with MspI for DNA from the WBS samples. The mean of abbreviations: C/N, C/N ratio; ECP, extracellular protein; ECPS, extracellular polysaccharide; TOC, total organic carbon; TN, total nitrogen. The number before the dash in the label represents the additional NH4 + -N content (0 (CK, ), 100 ( ), 300 (), 600 () and 1200 mg kg−1 (䊉)), whereas the number after the dash represents the sampling time. , the original WBS.

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Fig. 7. Relative abundance of the main methanotrophs in the WBS samples with NH4 + -N addition of 0 (CK), 100, 300, 600 and 1200 mg kg−1 . (a) Methylocaldum and/or Methylococcaceae; (b) Methylobacter; (c) Methylocystis and/or Methylosinus. Error bars denote the standard deviation of three replicates for the composite DNA of each treatment and sampling time point.

Relationship of the methanotrophic community and environmental variables The relationship between the methanotrophic communities and the six environmental variables, including the C/N ratio, ECP, ECPS, TOC, TN, NO3 − -N and NH4 + -N, was determined by performing RDA (Fig. 8). The first axis of the RDA explained 41.8% of the variation in the methanotrophic communities, and positively correlated with the C/N ratio and TOC. The NO3 − -N, NH4 + -N and TN contents were negatively related with the first axis. The second axis of the RDA explained 9.6% of the variation in the methanotrophic communities and this axis was positively related to ECP. The seven environmental variables explained 52.1% of the total variance, with the following order of importance (explicative variable) of the methanotrophic community data: NH4 + -N > C/N ratio > NO3 − N > TN > ECPS > TOC > ECP. The variance inflation factors of these six parameters were all less than 20, but only NH4 + -N and the C/N ratio had a significant influence on the variance by the Monte Carlo permutation test (P = 0.024 and 0.032, respectively). Discussion In this study, the addition of NH4 + -N ranging from 100 to 300 mg kg−1 could stimulate CH4 oxidation in WBS samples (Fig. 1),

although the NO3 − -N content of the original WBS was high (673 mg kg−1 ) at the first stage. The nitrogen source fixed by pure cultures of methanotrophs has been shown to be NO3 − -N rather than NH4 + -N [27,28]. However, in our previous study, it was also observed that NH4 + -N in WBS samples had a more stimulative effect for methanotroph activity than NO3 − -N at lower concentrations [42]. Further study on the mechanism of the stimulative effect of NH4 + -N on CH4 oxidation in WBS is therefore required. The addition of 600 mg kg−1 of NH4 + -N showed an inhibitory effect on CH4 oxidation in the first 4 days, but a stimulatory effect occurred between days 8 and 15, while the addition of 1200 mg kg−1 of NH4 + -N had a longer (∼8 days) inhibitory effect on CH4 oxidation. This indicated that the stimulatory or inhibitory effect of NH4 + -N addition on CH4 oxidation in WBS was largely dependent on NH4 + -N content. A higher inhibitory content of NH4 + -N on CH4 oxidation was observed in this study compared to other reports in which the NH4 + -N contents of landfill cover soils were amended to 14–40 mg kg−1 [4,33]. This suggested that besides NH4 + -N contents, the effect of NH4 + -N on CH4 oxidation in soils depended on many factors, such as the species of N, CH4 concentration, pH and the methanotrophic community in the soils [34]. Since ammonia is a competitive inhibitor of MMO [2,37,38], a high CH4 concentration can relieve the inhibitory effect of NH4 + -N on CH4 oxidation [42]. The inhibition of NH4 + on CH4 oxidation is also dependent on pH because NH3 is the active substrate for MMO, and it is more toxic than NH4 + [28,37,38]. At the end of the experiment, the pH value of the WBS samples ranging from 7.24 to 7.86 indicated that pH was not the main factor influencing the inhibitory effect of NH4 + -N on CH4 oxidation. Although the C/N ratio and TN content ranged from 28 mg kg−1 to 67 mg kg−1 and 499 mg kg−1 to 1206 mg kg−1 , respectively, on day 15, the CH4 oxidation rate showed a rapid decrease in the experimental treatments after day 22, and remained stable during the period between days 29 and 56. It has been reported that the CH4 oxidation rates have often exhibited a peak followed

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by a decrease to a lower steady state value in long-term laboratory column experiments simulating landfill cover or biofilter environments [20,35,41,43]. It has been hypothesized that the accumulations of EPS caused the decrease in CH4 oxidation rate due to clogging of soil pores, which led to short-circuiting of LFG through the soil material and/or impeded gas diffusion, thus reducing transfer of substrates into the cells [15,41,43]. The increase in the ECPS and ECP contents of the WBS samples from days 15 to 30 might explain the rapid decrease in the CH4 oxidation rate. In the last stage the ECP contents continued to increase, and reached 281–559 mg kg−1 on day 56, while the ECPS content decreased a little in all the experimental treatments, indicating that, compared to ECP, ECPS was more easily used as a carbon source for cell synthesis under conditions of nutrient shortage. EPS accumulation might be an adaptive strategy for methanotrophs and other bacteria for surviving in environments where these nutrients are imbalanced and there is O2 deficiency [32,34]. Type I methanotrophs Methylocaldum and Methylobacter, and type II methanotrophs (Methylocystis and Methylosinus) were abundant in the WBS samples. The type I methanotroph Methylocaldum dominated in the original WBS sample, which might be due to the high nutrient condition such as TOC (6.0%, w/w). In previous studies, type I methanotrophs have been reported to outcompete type II methanotrophs in high-nutrient, high-oxygen environments [3,19,40]. After exposure to LFG, the abundance of Methylobacter increased in all the experimental treatments, but a higher abundance of Methylobacter occurred in the WN300, WN600 and WN1200 treatments compared to those in the CK and WN100 treatments. This indicated that the relatively higher content of NH4 + -N addition (300–1200 mg kg−1 ) could lead to Methylobacter predominating in the WBS samples. This result is in agreement with other studies indicating that Methylobacter dominated in Nrich environments [13,30]. The peak abundance of Methylobacter occurred in the WN1200 treatment on day 30, which was later than those in the WN100, WN300 and WN600 treatments on day 15. This is likely to be due to the fact that an NH4 + -N addition of 1200 mg kg−1 inhibited the growth of Methylobacter in the first stage of activity. Compared to type I methanotrophs, the abundance of the type II methanotroph Methylocystis was low (0.2%) in the original WBS sample. However, after day 30, the abundance of Methylocystis showed an increasing trend, and reached 2.7–16.9% in the CK, WN100, WN300 and WN600 treatments, while it was close to 0 in the WN1200 treatment. This might be because, except for the WN1200 treatment, the N-limiting condition (the low NH4 + -N contents of 0.2–70.2 mg kg−1 ) occurred in the experimental treatments between days 30 and 56, leading to the growth of type II methanotrophs. Although the relative abundance of methanotrophs in the clone libraries of pmoA for DNA of the soil samples on day 56 was a little different from that in the T-RFLP profile due to sequences lower than 50, it also confirmed that Methylosinus dominated in the N-limiting condition. A similar result was obtained by De Visscher and Van Cleemput [9], who showed that type I methanotrophs grew faster than type II when inorganic N was not limiting in the first phase, but after a steady-state of a few weeks a new growth phase was probably dominated by N2 -fixing type II methanotrophs. In conclusion, the addition of NH4 + -N ranging from 100 to 300 mg kg−1 could stimulate CH4 oxidation in WBS samples at the first stage, while the addition of NH4 + -N contents of 600 and 1200 mg kg−1 had an inhibitory effect on CH4 oxidation. The community and activity of methanotrophs in the WBS samples varied with NH4 + -N content and incubation time. The NH4 + -N content and C/N ratio had a significant influence on the methanotrophic community in the WBS. These findings will be helpful for understanding the role of biotic and abiotic factors affecting the diversity and activity of methanotrophs in landfill cover soils,

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Acknowledgments This work was financially supported by the National Natural Science Foundation of China with Grant Nos. 41001148, 51178411 and 41371012, as well as the Zhejiang Province Natural Science Foundation for Distinguished Young Scholars (LR13E080002).

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