Effects of trichloroethylene on community structure and activity of methanotrophs in landfill cover soils

Soil Biology & Biochemistry 78 (2014) 118e127

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Effects of trichloroethylene on community structure and activity of methanotrophs in landfill cover soils Jiao-Yan Kong, Yun Bai, Yao Su, Yijun Yao, Ruo He* Department of Environmental Engineering, Zhejiang University, Hangzhou 310058, China

a r t i c l e i n f o

a b s t r a c t

Article history: Received 19 March 2014 Received in revised form 23 July 2014 Accepted 26 July 2014 Available online 13 August 2014

Landfill cover soil (LCS) plays an important role in mitigating the emission of CH4 and volatile organic gases from landfills to the atmosphere. In this study, effect of trichloroethylene (TCE) on community structure and activity of methanotrophs, as well as TCE degradation efficiency was investigated in waste biocover soil (WBS), which was collected from a landfill bioreactor treating organic waste, in comparison with LCS. The CH4 oxidation activity and TCE degradation rate were higher in WBS compared to those in LCS. The TCE degradation rates in both soils were enhanced with the increase of TCE concentration within 50 ppmv. Compared to LCS, the TCE inhibitory concentration that caused inhibition of CH4 oxidation activity was greater for WBS. The abundance of mmoX was similar in both soils during the whole experiment, while the average abundance of pmoA in WBS was about two orders of magnitudes higher than in LCS. Type I methanotrophs (Methylocaldum, Methylomonas, Methylosarcina and Methylobacter) and type II methanotrophs (Methylocystis) were abundant in both soils. Among them, type I methanotrophs Methylocaldum and Methylobacter dominated in WBS, while type II methanotrophs Methylocystis predominated in LCS. The relative abundance of Methylobacter increased with an increase of TCE concentration and exposure time in both soils, especially in WBS, indicating that Methylobacter seemed tolerant to TCE and/or may play an important role in the TCE degradation. © 2014 Published by Elsevier Ltd.

Keywords: CH4 oxidation Methanotrophs Trichloroethylene Landfill gas Landfill cover soil

1. Introduction Landfill gas (LFG) mainly originates from the biodegradation of organic matter in landfills, which consists of methane (CH4) (55e60%, v/v), carbon dioxide (CO2) (40e45%, v/v) and trace gases such as halogenated, aromatic hydrocarbons and sulfur-containing compounds (Scheutz and Kjeldsen, 2004). Both CH4 and CO2 are important greenhouse gases. The global warming potential of CH4 is about 25 times higher than that of CO2 on a 100-year time frame (IPCC, 2007). Landfills, as one of the major anthropogenic sources of CH4 emissions, account for 6e12% of the total global CH4 emissions. It is estimated that 35e69 Tg CH4 is released annually into the atmosphere from landfills (IPCC, 2007). Although the concentration of trace gases is less than 2% (v/v) in LFG, many trace gases such as toluene and trichloroethylene (TCE) are on the United State Environmental Protection Agency priority pollutant list and pose a potential serious risk to public health (Assmuth and Kalevi, 1992).

* Corresponding author. Tel./fax: þ86 571 88982221. E-mail address: [email protected] (R. He). http://dx.doi.org/10.1016/j.soilbio.2014.07.018 0038-0717/© 2014 Published by Elsevier Ltd.

Landfill cover soil (LCS) is the environmental interface between deposited waste and the atmosphere, which can serve as a biofilter to reduce the emission of LFG pollutants. It has been reported that 6e100% of CH4 that escapes from landfills is consumed by LCS €rjesson et al., 2004; Einola et al., 2007; (Barlaz et al., 2004; Bo Scheutz et al., 2009). Aerobic methanotrophs are the primary mediators of CH4 consumption in oxic layers of LCS before CH4 releases into the atmosphere (Henneberger et al., 2012). Novel thermoacidophilic aerobic methanotrophs within the Verrucomicrobia were recently discovered in geothermal areas (Dunfield et al., 2007; Pol et al., 2007; Islam et al., 2008). Aerobic methanotrophs mainly belong to the Proteobacteria and can be divided into two taxonomic groups: type I (belonging to the Methylococcaceae family of gProteobacteria) and type II methanotrophs (including the genera Methylocystis, Methylosinus, Methylocella and Methylocapsa in aProteobacteria), based on their cell morphology, ultra-structure, phylogeny and metabolic pathways (Hanson and Hanson, 1996; Semrau et al., 2010). CH4 is oxidized to methanol by methane monooxygenase (MMO), which exists in two forms: the membrane bound particulate MMO (pMMO) and the cytoplasmic soluble MMO (sMMO) (Hanson and Hanson, 1996; Semrau et al., 2010). pMMO is known to be present in all known methanotrophs except

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for the genera Methylocella (Dedysh et al., 2000) and Methyloferula (Vorobev et al., 2011), whereas sMMO appears to occur only in certain methanotroph strains. MMO, especially sMMO, has a broad substrate specificity and can oxidize a range of recalcitrant hydrocarbons, including TCE, ethanes and ethenes (Semrau et al., 2010). It has been reported that Methylocystis sp. strain M, Methylocystis strain SB2, Methylomonas sp. strain MM2, Methylomonas methanica 68-1, Methylomicrobium album BG8, Methylosinus trichosporium OB3b and methanotrophic culture can simultaneously cometabolize halogenated and aromatic hydrocarbons in the presence of CH4 (Oldenhuis et al., 1989; Koh et al., 1993; Vlieg et al., 1996; Han et al., 1999; Shukla et al., 2009, 2010; Im and Semrau, 2011). Under lowcopper growth conditions, Methylosinus trichosporium OB3b can degrade TCE at the biotransformation capacity of ~0.25 mg mg1 dry weight of cell mass (Taylor et al., 1993). CH4 oxidation rates vary with physical and chemical characteristics of landfill covers, such as soil texture, particle composition, water content, organic carbon content, temperature and nutrient (Hanson and Hanson, 1996; Scheutz and Kjeldsen, 2004; Scheutz et al., 2009; Wang et al., 2011). In addition to the capacity of CH4 oxidation, LCS has a significant potential for degradation and cometabolic degradation of volatile organics such as vinyl chloride (VC) and TCE (Scheutz et al., 2004). However, MMO-mediated degradation of chlorinated hydrocarbons can generate toxic intermediates that may inactivate cells. For examples, oxidation of TCE by MMO will produce TCE-epoxide (Little et al., 1988). Some of hydrolysis products of TCE-epoxide can covalently bind to MMO, which may result in the death of cells (Fox et al., 1990; Nakajima et al., 1992). In addition, there is competition between CH4 and the chlorinated co-substrate for the (s)MMO active site (Smith and Dalton, 2004). Therefore, understanding the effect of trace gases in LFG such as TCE on CH4 biological process would be helpful to control the emission of LFG pollutants into the atmosphere. Biocover soils, such as compost, waste biocover soil (WBS) and mineralized waste, have been demonstrated to have a high CH4 oxidation due to their well-distributed particle size, high organic matter content and active microorganism activity (Scheutz and Kjeldsen, 2004; Bogner et al., 2010; He et al., 2012a). However, little is known about the effect of halogenated compounds on CH4 oxidation and methanotrophic community in biocover soils. In this study, TCE was chosen as the target compound to evaluate the CH4 oxidation potential and TCE degradation efficiency under varied TCE concentrations in WBS compared with LCS. The CH4 oxidation activity and biodegradation rate of TCE were tracked in microcosms at a series of TCE concentrations (0, 0.5, 5, 20 and 50 ppmv) representing reported TCE concentrations at landfills that range from 0.001 to 20 ppmv (Chiemchaisri et al., 2001; Shafi et al., 2006). Genes pmoA and mmoX, respectively, encoding a key subunit of pMMO and sMMO, were used to analyze the identity and diversity of methanotrophs in both soils using quantitative PCR (Q-PCR), terminal restriction fragment length polymorphism (T-RFLP) and cloning. It was hypothesized that methanotrophs in both soils would have different CH4 oxidation activity and present different response to the TCE addition. Methanotrophs dominating in the soil samples exposed to high concentrations of TCE might be tolerant to TCE and play an important role in the TCE degradation.

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bioreactor was described by Wang et al. (2011). The landfill bioreactor had been operated for ~2 years before taking the stabilized waste (WBS). LCS was collected at the 10e20 cm depth of cover soil from Dawuao landfill, which is located in Dawuao Mountain in Pingshui Town, Zhejiang Province. The sampling site located in an area where LCS had been in place for ~12 yr. After removing plants and large particles, the soils were air-dried and sieved through a 4 mm mesh. The physical and chemical properties of WBS were described previously (Wang et al., 2011; He et al., 2012a). The particle composition of the WBS was 61% of 2e4 mm, 33% of 0.02e2 mm and 6% of <0.02 mm. The pH was 7.7. The organic matter and total nitrogen contents of the WBS were 31 g kg1 and 1.3 g kg1, respectively. The particle composition of the LCS was 42.3% of 2e4 mm, 46.9% of 0.5e2 mm and 10.8% of 0.5 mm. The pH was 6.8. The organic matter and total nitrogen contents of the LCS were 17 g kg1 and 0.4 g kg1, respectively. 2.2. Landfill cover microcosms Approximately 50 g of the air-dried experimental soil was placed into a 400 ml serum bottle. The WBS sample was adjusted to the water content of 45% (w/w), at which the WBS was reported to have the highest CH4 oxidation activity (Wang et al., 2011). Because of its low water holding capacity (i.e. the saturated water content of LCS is about 30% (w/w)), it is impossible to adjust the LCS to the same water content as the WBS. Thus, the LCS sample was adjusted to the water content of 20% (w/w), at which the LCS sample was detected to have the highest CH4 oxidation capacity. The serum bottles were covered with cling film and allowed to equilibrate the soil water contents overnight (~12 h) at 30  C, and then sealed with butyl rubber stoppers. A certain volume of air was withdrawn from the serum bottle prior to injecting simulated LFG. Simulated LFG (CH4:CO2 ¼ 1:1) was injected into the serum bottles to obtain the CH4 and CO2 concentrations of 10% (v/v). High purity O2 was injected into the serum bottles to keep the O2 concentration at ~21% (v/v). Gas samples were withdrawn periodically from the headspace of the serum bottles for measuring the concentrations of CH4, CO2 and O2. The serum bottles were flushed with air and the initial concentrations of CH4, CO2 and O2 were resupplied to re-establish the initial gases concentrations each day. After one month of pre-incubation with simulated LFG, the WBS and LCS were denoted as the original material (0 d) for the experiment. The organic matter and total nitrogen contents were 17.7 and 0.6 g kg1 for the original WBS, and 8.2 and 0.3 g kg1 for the original LCS, respectively. Then TCE was injected into the serum bottles to achieve the concentrations of 0, 0.5, 5, 20 and 50 ppmv, respectively. The treatment with sterilized soils and NaN3 (0.13 mg g1 (dry weight)) were used as non-microbial controls to account for such things as adsorption. All treatments were performed in triplicate, and incubated at 30  C. Gas samples were withdrawn periodically from the headspace of the serum bottles for measuring CH4, CO2, O2 and TCE concentrations. The serum bottles were flushed with air, and then the initial concentrations of CH4, CO2, O2 and TCE were re-established in the fume hood as described above each day. The whole experiment lasted 54 days. 2.3. Sampling and analysis

2. Material and methods 2.1. Experimental soils Two types of soils, namely WBS and LCS, were used in this study. The WBS was collected from a landfill bioreactor (2 m3) treating organic waste with leachate recycle located in a village in Xindeng town, Zhejiang Province. The raw solid waste composition in the

Gas samples (100 ml) were periodically taken from the headspace of the serum bottles for measuring CH4 concentrations as described previously (Wang et al., 2011). The CH4 oxidation activity was calculated by applying zero-order kinetics as described by Kightley et al. (1995), and expressed by the mass of the CH4 oxidized per dried soil per hour (mg g1 h1). Gas samples (500 ml) were withdrawn from the headspace of the serum bottles for the

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measurement of TCE concentration, which was performed using a gas chromatograph (GC) equipped with a flame ionization detector with a SE-54 column (30 m  0.32 mm  1.5 mm). N2 was used as carrier gas at a rate of 15 ml min1. The temperatures of the oven, injector and detector were 90, 200 and 250  C, respectively. The detection limit of GC for TCE was ~0.4 ppmv. The treatment with the TCE concentration of 0.5 ppmv was carried out to investigate the effect of low concentrations of TCE on the activity and community of methanotrophs, despite that the variation of TCE concentration could not be measured in this study. The TCE degradation amount equaled the amount of TCE loss in the non-microbial control subtracted from that in the corresponding TCE concentration treatment. The TCE degradation amount presented a strong linear relationship with time within the first 6 h when the TCE concentrations in the headspace of the serum vials were higher than 20% of the initial injected TCE concentrations in this study. Therefore, the TCE degradation rate in this study was calculated by zero-order kinetics within about 5 h after TCE injection and normalized to the dry soil mass as described by Scheutz et al. (2004). 2.4. DNA extraction and Q-PCR analysis DNA extraction was performed with the E.Z.N.A.™ Soil DNA Kit (Omega Bio-Tek, Inc., Norcross, USA). Three replicates of each treatment and sampling time point were used and the extracted DNA was quantified using the Nanodrop ND-1000 spectrophotometer. The DNA from each replicate was mixed in equal amounts to compose the DNA of each treatment and sampling time point for Q-PCR and T-RFLP analysis. pmoA was amplified using the primers A189F (Holmes et al., 1995) and mb661R (Costello and Lidstrom, 1999). mmoX was amplified using the primer set of mmoX1 and mmoX2 (Miguez et al., 1997). Q-PCR was performed in triplicate of 15 ml of reaction mixture containing All-in-one qPCR Mix (GeneCopoeia, Inc., USA), 4.0 pmol of each primer and 1 ml template. Thermal cycler conditions of pmoA were as follows: an initial stage at 95  C for 10 min; 40 cycles of 95  C for 10 s, 58  C for 30 s, and 72  C for 30 s. Thermal cycler conditions of mmoX were as follows: an initial stage at 95  C for 10 min; 40 cycles of 95  C for 10 s, 56  C for 20 s, and 72  C for 30 s. Standards were made from 10-fold dilutions of linearized plasmids containing the same fragment of pmoA or mmoX that was cloned from amplified Methylosinus sporium DNA. A melting point analysis was conducted to confirm that no unspecific PCR products were generated and analyzed. The detection limit of Q-PCR was ~102 copies per reaction for pmoA and mmoX. The abundance of pmoA and mmoX was expressed as copies g1 dry weight soil. 2.5. T-RFLP analysis PCR for T-RFLP analysis was performed for pmoA using 50 6carboxyfluorescien-labeled forward primer A189F and unlabeled reverse primer mb661R as described by Kong et al. (2013). PCR products were purified using 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 as the procedure described by He et al. (2012b). Pellets were resuspended in 1.0 ml H2O, 0.5 ml of GeneScan™ 500LIZ™ Size Standard (Applied Biosystems, Foster City, CA, USA) and 9.5 ml of deionized formamide, 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

using the GeneMapper software system (Applied Biosystems, Foster City, CA, USA). The peak height threshold for T-RFLPs was set at 50 fluorescence units. Terminal restriction fragments (T-RFs) below 50 base pair (bp) were eliminated from all datasets. The remaining peaks were normalized, and their heights represented a percentage of the total peak height in each sample. Different methanotrophic communities were then subjected to a multivariate cluster analysis using BrayeCurtis similarity index with the open source statistical application PAST (Hammer et al., 2001). 2.6. Cloning and phylogenetic analyses DNA extracted from the sixteen soil samples (the three replicates of each treatment and sampling time point were used for DNA extraction) was quantified using the Nanodrop ND-1000 spectrophotometer. DNA extracted from the three replicates was mixed in equal amounts to be used as the template for PCR amplification of each treatment and sampling time point. The PCR reaction mixture was described by Kong et al. (2013) with the primer sets of A189F/ mb661R for pmoA and mmoX1/mmoX2 for mmoX, respectively. Thermal cycler conditions were as follows: denaturation at 94  C for 3 min; 30 cycles of 94  C for 30 s, 58  C for pmoA or 56  C for mmoX for 30 s, and 72  C for 40 s; and a final extension step of 72  C for 10 min. PCR product purification and cloning was carried out as described previously (Kong et al., 2013). T-RFLP analysis of individual pmoA clone was carried out to check for differences in T-RF size between in silico predicted and experimentally determined peaks. Raw pmoA and mmoX sequences were trimmed with primer sequences removed using DNASTAR software (DNASTAR Inc. Madison, USA), and then inspected for chimeras by searching for large sequence regions of unexpected nucleotide changes when compared with the reference sequences. Sequences were aligned with related sequences extracted from GenBank and translated to get deduced amino acid sequences. Phylogenetic tree was constructed using MEGA 5 software applying the maximum-likelihood method as described by He et al. (2012b). Sequences in this study have been submitted to the GenBank database under accession numbers KF055930eKF055953 and KJ561912eKJ561928. 3. Results 3.1. CH4 oxidation activity After incubation with simulated LFG for a month, the CH4 oxidation activities in LCS and WBS were 15 and 210 mg g1 h1, respectively (Fig. 1). In WBS, there was no obvious difference in the CH4 oxidation activity among the treatments with the varied TCE concentrations during the first 15 days (P > 0.6). The CH4 oxidation activity in WBS presented an increasing trend between days 22 and 42. After day 42, the CH4 oxidation activity in WBS fluctuated in a range of 690e811 mg g1 h1 until the end of the experiment except for the treatment exposed to the TCE concentration of 50 ppmv. The CH4 oxidation activity was not significantly affected by the exposure of TCE concentrations ranging from 0.5 to 50 ppmv at the first 42 days. However, from day 48, a significant decrease was observed in the CH4 oxidation activity at the TCE concentration of 50 ppmv (P < 0.01). After exposure to LFG, the CH4 oxidation activity was enhanced in all experimental LCS treatments and reached 90e170 mg g1 h1 on day 36, which was 6e12 times higher than that of the original (day 0). After day 36, there was a decreasing trend in the CH4 oxidation activity of LCS, and it dropped to 35e102 mg g1 h1 on

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8 15 22 30 36 42 48 54 Time (d) 0 ppmv 0.5 ppmv 5 ppmv 20 ppmv 50 ppmv

Fig. 1. CH4 oxidation rates of soils (dry weight) in the presence of TCE over time. Sign in Figure means the initial TCE concentrations.

day 42. Among the experimental treatments with varied TCE concentrations (i.e. 0, 0.5, 5, 20 and 50 ppmv), the highest CH4 oxidation activity was observed in the LCS sample exposed to the TCE concentration of 0.5 ppmv. Compared to the treatment without TCE addition, the CH4 oxidation activity was lower in the treatments with the TCE concentrations ranging from 5 to 50 ppmv.

3.2. TCE biodegradation rate After exposure to LFG containing TCE, the TCE degradation rate of WBS showed an increasing trend over time during the first 22 days, and reached 0.083, 0.162 and 0.364 mg g1 h1 at the TCE concentrations of 5, 20 and 50 ppmv, respectively, on day 22 (Fig. 2). After that (day 22), the TCE degradation rate of WBS fluctuated in ranges of 0.047e0.103, 0.119e0.201 and 0.269e0.379 mg g1 h1 at the TCE concentrations of 5, 20 and 50 ppmv, respectively, until the end of the experiment. The TCE degradation rates of LCS at the concentrations of 5, 20 and 50 ppmv were 0.014, 0.032 and 0.072 mg g1 h1, respectively, on day 0. After exposure to LFG containing TCE, the trend of the TCE degradation rate in LCS was similar to that in WBS. Compared to WBS, the TCE degradation rate was lower in LCS, only 56% of that in WBS on average.

3.3. Abundance of pmoA Q-PCR analysis showed that the abundance of pmoA varied with soil types, TCE concentrations and exposure time (Fig. 3). The abundance of pmoA was 8.9  108 and 5.7  107 copies g1 in the original WBS and LCS (day 0), respectively. After exposure to LFG, the abundance of pmoA in WBS increased to 1.7  109e3.6  109 copies g1 on day 15, and then fluctuated within a range of 0.9  109e2.6  109 copies g1 between days 30 and 54.

0.6 0.5

0.4 0.3 0.2 0.1 0.0 1

8 15 22 30 36 42 48 54 Time (d) 5 ppmv 20 ppmv 50 ppmv

Fig. 2. TCE biodegradation rates of soils (dry weight) over time. Sign in Figure means the initial TCE concentrations.

After exposure to LFG for 15 days, the abundance of pmoA in LCS increased slightly and the highest abundance of pmoA was observed in the LCS sample exposed to the TCE concentration of 0.5 ppmv, followed by the treatment without TCE addition. There was no significant difference in the abundance of pmoA between the LCS samples exposed to the TCE concentrations of 20 and 50 ppmv on day 15 (P ¼ 0.14). The abundance of pmoA in all the LCS samples decreased slightly on day 30, and remained stable until the end of the experiment. Compared to WBS, the average abundance of pmoA in LCS was about two orders of magnitude lower during the whole experiment. 3.4. Abundance of mmoX The abundance of mmoX was below the detection limit in the original WBS (0 d), but it was detected in the WBS samples with the abundance of 1.4  106e5.6  106 copies g1 after exposure to TCE for 15 days (Fig. 4). On day 30, the mmoX was also detected in the WBS sample without TCE addition with the abundance of 5.6  105 copies g1. The abundance of mmoX present an increasing trend in the WBS samples with the increase of TCE concentrations on day 30. However, on day 54, the abundance of mmoX decreased to 6.9  105 copies g1 in the WBS with the TCE addition of 50 ppmv, which was near to that without TCE addition. The abundance of mmoX was 2.5  105 copies g1 in the original LCS sample. After exposure to TCE and LFG, the abundance of mmoX increased in all the LCS samples on day 15. On day 30, the abundance of mmoX decreased to 9.3  105e3.3  106 copies g1 in the LCS samples. However, the abundance of mmoX increased a little at the end of the experiment (day 54). No significant difference was observed in the abundance of mmoX among all the LCS samples without TCE addition on days 15, 30 and 54 (P > 0.75).

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Time (d)-TCE concentration (ppmv) Fig. 3. Quantitative PCR (Q-PCR) of pmoA for DNA from soil (dry weight) samples on day 15, 30 and 54 as well as the original soil (0 d). The number before the dash in the label represents the sampling time (original (0 d), days 15, 30 and 54); the number after the dash represents the initial TCE concentration (ppmv).

3.5. Characterization of methanotrophic community To assess the overall community structure of methanotrophs, DNA extracted from the 16 soil samples for the T-RFLP analysis of each soil was combined together and used to create clone libraries of pmoA and mmoX for each soil. A total of 288 and 75 pmoA and mmoX clones, respectively, were obtained from DNA samples of WBS and the sequences were subjected to BLASTn searches using GenBank. Among 288 pmoA clones obtained from WBS, 261 clones (90.6%) showed 92e100% sequence similarities to Methylocaldum sp. S8 (AB900159), Methylocaldum sp. 5FB (AJ868403) and Methylocaldum tepidum (MTU89304); 22 clones (7.6%) showed 91% similarity to Methylobacter sp. BB5.1 (AF016982); and 5 clones (1.7%) had 99% similarity to Methylocystis sp. KS3 (AJ459033) (Fig. 5). All the mmoX clones obtained from WBS showed 99e100% sequence similarities to Methylocaldum sp. S8 (AB900160), uncultured bacterium (EF472923) and uncultured bacterium (EF472928) (Fig. 6). Among 52 pmoA clones obtained from DNA from the LCS, 46 clones (88.5%) had 97e99% similarities to Methylocystis spp. (i.e. Methylocystis sp. M212 (JN036528), Methylocystis sp. 62/12 (AJ459003), Methylocystis sp. LW5 (AF150791), Methylocystis sp. SS2C (AB636307), Methylocystis sp. m231 (DQ852353), Methylocystis sp. 5FB1 (AJ868406) and Methylocystis sp. 39 (AJ459045)), 2 clones (3.8%) showed 84% similarity to Methylovulum miyakonense HT12 (AB501285), one clone (1.9%) had 91% similarity to Methylobacter sp. BB5.1 (AF016982); one clone (1.9%) had 94% similarity to Methylosarcina lacus (AY525413); one clone (1.9%) had 85%

Fig. 4. Quantitative PCR (Q-PCR) of mmoX for DNA from soil (dry weight) samples on day 15, 30 and 54 as well as the original soil (0 d). The number before the dash in the label represents the sampling time (original (0 d), days 15, 30 and 54); the number after the dash represents the initial TCE concentration (ppmv).

similarity to Methylococcus capsulatus (AF533666); and one clone (1.9%) had 97% similarity to uncultured bacterium (EU193280). Among 67 mmoX clones obtained from DNA from the LCS, 39 clones (58.2%) showed 94e99% similarities to Methylocystis sp. 41 (AJ458532), Methylocystis sp. M (MSU81594), Methylocystis sp. 5FB2 (AJ868419), Methylocystis sp. S284 (HE798550); and 28 clones (41.8%) had 94% and 90% similarity to uncultured bacterium(EU131087) and Methylomonas sp. LW13 (AY007290), respectively.

3.6. Methanotrophic diversity and abundance with TCE concentrations To compare the effect of TCE concentrations on methanotrophic community, pmoA-based T-RFLP profiles were conducted for the original soil (day 0) and the soil samples exposed to LFG containing various concentrations of TCE on days 15, 30 and 54. 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 in WBS, while it was related with Methylococcus and/or Methylocaldum in LCS; the 244-bp T-RF was related with Methylocystis; the 437-bp T-RF was related with Methylosarcina; the 508bp T-RF was related with Methylobacter (Fig. 5). The T-RFLP analysis of pmoA clones showed that the deviation between the predicted by in silico digestion and those experimentally determined T-RF sizes was within 3 bp.

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L1A6(4)-79bp 52 L1E3(2)-79bp 99 Methylocaldum sp. S8(AB900159) 85 L2D8(12)-79bp 98 L2F9(19)-79bp L2D4(3)-79bp

Methylocaldum sp. 5FB(AJ868403)

78 95 L2E8(11)-79bp 98 74 Methylocaldum tepidum (MTU89304) 97 Methylocaldum sp. 05J-I-7 (EU275141) 100 Methylocaldum sp. ML100 (AF510078) L1D6(205)-79bp 98 L1H4(5)-79bp T4A(1)-79bp 84

Methylosoma sp. TFB (GQ130273) Methylococcaceae bacterium OS501 (AB636304) Methylococcus capsulatus (AF533666)

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Methylosarcina lacus (AY525413) Methylomonas sp. ML64 (AF510080) Methylococcaceae bacterium M200 (HM564018) Methylovulum miyakonense (AB501285) Methylobacter sp. LW12 (AY007285)

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Methylobacter sp. BB5.1 (AF016982) T9A(2)-508bp

Methylosinus sp. ML19 (AF510081) Methylosinus sp. B4S (AB683146)

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T2C(2)-244bp

Methylocystis sp. M212 (JN036528)

100 L2G4(5)-244bp

Methylocystis sp. KS3(AJ459033)

86 99 91 69

T10E(1)-244bp

Methylocystis sp. LW5 (AF150791)

T6A(1)-244bp 96 T1G(2)-244bp

Methylocystis sp. 39 (AJ459045)

T6H(21)-244bp

Type II Methanotrophs

68 Methylocystis sp. 62/12 (AJ459003) T11H(12)-244bp 66 Methylocystis sp. SS2C (AB636307) T1F(1)-244bp

Methylocystis sp. m231 (DQ852353) Methylocystis echinoides (AJ459039) T3G(4)-244bp T2A(2)-244bp

Methylocystis sp. 5FB1 (AJ868406) Nitrosomonas europaea (AF037017) 0.1 Fig. 5. Maximum-likelihood tree of deduced pmoA sequences. The scale bar represents 0.1 substitutions per nucleotide position. Bootstrap values >50 are shown. The meanings of the first letter L and T in the clone names denote the cloning sequence of pmoA for DNA from WBS and LCS, respectively. The number of clones assigned to each operational taxonomic unit (OTU) by restriction fragment length polymorphism (RFLP) is shown in parentheses. Predicted T-RF size (bp) determined by in silico digestion with Msp I is listed.

Type I methanotrophs (Methylocaldum, Methylosarcina and Methylobacter) and type II methanotrophs (Methylocystis) were the main methanotrophs in WBS, accounting for 89.7e95.2% of the total T-RFs abundance (Fig. 7). Among them, type I methanotroph Methylocaldum dominated in the original WBS sample, with a relative abundance of 88.5%. After exposure to LFG, the relative abundance of Methylocaldum did not change much in the WBS samples without the TCE addition, ranging from 79.8 to 89.7% of the total T-RFs abundance. However, in the WBS samples exposed to TCE, there was a decreasing trend in the relative abundance of Methylocaldum with the increasing TCE concentration and exposure time. In the WBS samples exposed to the TCE concentration of 50 ppmv, the relative abundance of Methylocaldum dropped to 27.2% at the end of the experiment. Although Methylobacter was not predominant in the original WBS sample with a relative abundance

of 2.1%, it exhibited an increasing trend in the relative abundance in WBS with the increase of TCE concentration and exposure time. In the WBS samples exposed to the TCE concentration of 50 ppmv, the relative abundance of Methylobacter increased to 62.2% at the end of the experiment. Methylosarcina was identified in WBS, but the relative abundance of Methylosarcina was low within a range of 0e1.3%. Type II methanotroph Methylocystis was also identified in WBS with the relative abundance of 0.7e5.1% during the experiment. Type II methanotrophs Methylocystis dominated in the original LCS samples with a relative abundance of 78.1%. After exposure to LFG, the relative abundance of Methylocystis dropped a little in the LCS samples with the fluctuating relative abundance of 42.7e69.9%. The relative abundance of Methylobacter did not show remarkable difference among the LCS samples with TCE addition during the

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Fig. 6. Maximum-likelihood tree of deduced mmoX sequences. The scale bar represents 0.1 substitutions per nucleotide position. Bootstrap values >50 are shown. The meanings of the first letter L and T in the clone names denote the cloning sequence of pmoA for DNA from WBS and LCS, respectively. The number of clones assigned to each operational taxonomic unit (OTU) by restriction fragment length polymorphism (RFLP) is shown in parentheses.

first 30 days. However, a higher relative abundance of Methylobacter was observed in the LCS samples exposed to the TCE concentrations of 20 and 50 ppmv compared to that without TCE addition on day 54. The relative abundance of Methylococcus and/or Methylocaldum in the LCS samples increased to 16.9e35.0% between days 15 and 30. Methylosarcina was not found in the original LCS samples, but it was detected in the LCS samples after exposed to LFG with the relative abundance ranging from 1.7 to 10.5%. A multivariate cluster analysis of T-RFLP profiles using BrayeCurtis similarity index showed that the community structure of methanotrophs separated clearly with soil types, WBS and LCS (Fig. 8). In the WBS samples, the methanotrophic community clustered with the increase of TCE concentrations and exposure time. The WBS samples exposed to the higher TCE concentrations (20 and 50 ppmv) and longer time (day 54) grouped tightly. 4. Discussion The average abundance of pmoA in WBS was about two orders of magnitude higher than in LCS during the whole experiment, indicating that WBS was more favorable to the growth of methanotrophs and further led to a higher CH4 oxidation activity in WBS (Fig. 1). A decline in CH4 oxidation activity of LCS was observed after day 42, and then the activity remained stable until the end of the experiment. Similar results were observed in long-term laboratory column experiments simulating landfill cover or biofilter

environments (Streese and Stegmann, 2003; Scheutz and Kjeldsen, 2004; Wilshusen et al., 2004b). The extracellular polymeric substances contents of LCS presented an increasing trend at the last stage of the experiment (Fig. S1), which might lead to a decrease in the CH4 oxidation rate, because the clogging of soil pores could impede gas diffusion and thus reduce transfer of substrates into cells (Streese and Stegmann, 2003; Wilshusen et al., 2004a; Haubrichs and Widmann, 2006). However, further studies such as nutrient availability also need to be conducted to confirm the reason for the decrease in the CH4 oxidation activity of landfill covers. High concentrations of TCE addition (50 ppmv for WBS, 20 and 50 ppmv for LCS) had an obvious inhibitory effect on CH4 oxidation. A significant decrease in CH4 oxidation rates after exposure to TCE was also observed in methanotrophic cultures (Broholm et al., 1990; Alvarez-Cohen and McCarty, 1991). It was hypothesized that some short-lived, highly reactive intermediates, such as TCE epoxide, were responsible for the inhibitory effect on CH4 oxidation (Fox et al., 1990; Vlieg et al., 1996). Competitive inhibition might also have occurred in the presence of TCE during CH4 oxidation due to the crucial step of TCE degradation catalyzed by MMO, which is also the key enzyme in CH4 oxidation (Semprini et al., 1991). There was no obvious difference in the CH4 oxidation activity of the two soils exposed to the TCE concentrations of 0.5 and 5 ppmv in comparison with that without TCE addition. Oldenhuis et al. (1989) also reported that TCE was not toxic to methanotrophic cultures

J.-Y. Kong et al. / Soil Biology & Biochemistry 78 (2014) 118e127

Fig. 7. Relative abundance of terminal-restriction fragments (T-RFs) of pmoA amplicons. The number before the dash in the label represents the sampling time (original (0 d), days 15, 30 and 54); the number after the dash represents the initial TCE concentration (ppmv).

with a concentration below 15 mg l1. Compared to LCS, the inhibitory concentration and exposure time of TCE on CH4 oxidation was higher and longer in WBS, likely due to the higher abundance of methanotrophs in WBS. It indicated that WBS provided a good nutrient condition for the growth of methanotrophs and thus better be able to withstand toxicity of TCE and its intermediates. TCE can be biodegraded by a diversity of bacteria such as methanotrophs, and species of Pseudomonas (Pseudomonas putida and Pseudomonas cepacia) (Landa et al., 1994; Cox et al., 1998). In this study, the TCE degradation rate of WBS increased with time during the first 22 days, with a higher abundance of pmoA and mmoX on day 15. The presence of CH4 may have also induced a nonspecific oxygenase (MMO) that can also catalyze the degradation of TCE (Broholm et al., 1990; Alvarez-Cohen and McCarty, 1991; McFarland et al., 1992). Similar result was obtained by Wilson and Wilson (1985) in the study of the methanotrophic biodegradation of TCE in sand columns where indigenous methanotrophic microorganisms were biostimulated by feeding natural gas. In this study, the TCE degradation rate increased with TCE concentration in both soils as observed by Smith et al. (1997). Type I methanotrophs (Methylocaldum, Methylomonas, Methylosarcina and Methylobacter) and type II methanotrophs (Methylocystis) were abundant in the two soils. Among them, type I methanotroph Methylocaldum dominated in WBS, while type II methanotrophs Methylocystis dominated in LCS (Figs. 5 and 6). Type II methanotrophs are generally more prevalent in areas with low O2 and high CH4 concentrations, while type I methanotrophs prefer

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inverse conditions (Amaral and Knowles, 1995; Henckel et al., 2000; Stralis-Pavese et al., 2004; Henneberger et al., 2012). The result obtained by Knief et al. (2006) also demonstrated that type I methanotrophs became active at CH4 mixing ratios as high as 500 ppmv. In addition to CH4 and O2 concentrations, methanotrophic community also changes with other environmental factors such as nitrogen availability, nutrient condition, pH value (Hanson and Hanson, 1996; Bodelier and Laanbroek, 2004; Scheutz et al., 2009). In this study, type I methanotrophs were more abundant than type II methanotrophs in WBS, which might be attributed to the high nutrient condition such as organic content (3.1%, w/w). In previous studies, type I methanotrophs have been reported to outcompete type II methanotrophs in high-nutrient and highoxygen environment (Henckel et al., 2000; Stralis-Pavese et al., 2004; Henneberger et al., 2012). The abundance of mmoX was below the detection limit in the original WBS (0 d), but it was detected in the WBS samples after exposure to TCE for 15 days, indicating that TCE exposure at the concentration within 50 ppmv could stimulate the growth of mmoX-based methanotrophs, such as Methylocaldum sp. S8. But a low abundance of mmoX in the WBS sample exposed to the TCE concentration of 50 ppmv on day 54 suggested that the longer exposure of higher concentrations of TCE would be toxic for mmoXbased methanotrophs in WBS. Similar result was observed in the abundance of pmoA and mmoX in LCS. During the experiment, the abundance of mmoX in the WBS samples exposed to various TCE concentrations was similar to that in LCS (105e106 copies g1), but a higher TCE degradation rate occurred in WBS than in LCS. It might be attributed to the high abundance of methanotrophs possessing pMMO in WBS (two orders of magnitude higher than that in LCS during the whole experiment), which exhibit a certain TCE degradation rates ranging from negligible to rates comparable to sMMO (Anderson and McCarty, 1997; Lontoh and Semrau, 1998; Im and Semrau, 2011). A decreasing trend in the relative abundance of Methylocaldum with the increase of TCE concentrations and exposure time was observed in WBS, suggesting that most Methylocaldum members were sensitive to TCE. This has also been observed in methanotrophic enrichment, where Methylocaldum sp. (Ib559) was not detected when the consortium was exposed to TCE (Choi et al., 2013). The abundance of Methylobacter increased from 2.1% in the original WBS samples to 62.2% in the sample exposed to the TCE concentration of 50 ppmv on day 54. A higher abundance of Methylobacter was also observed in the LCS samples exposed to the TCE concentration of 20 and 50 ppmv than those exposed to the TCE concentration of 0, 0.5 and 5 ppmv at the end of the experiment. This suggested that Methylobacter might be tolerant to TCE and/or play an important role in the TCE degradation in both soils, especially in WBS. This was in agreement with Choi et al. (2013) that the relative abundance of Methylobacter in methanotrophic enrichment increased after the TCE addition. Moreover, Methylobacter sp. BB5.1, a bacteria of high similarity with many of the pMMOcloned sequences in this study, has been reported to exhibit TCE degradation activity when the cells are incubated in a medium containing copper (Smith et al., 1997). In conclusion, WBS was more favorable to the growth of methanotrophs than LCS. A higher CH4 oxidation activity and a higher TCE degradation rate occurred in WBS compared to those in LCS. Activity and community of methanotrophs in both soils varied with TCE concentrations and exposure time. Type I methanotrophs (Methylocaldum and Methylobacter) dominated in WBS, while type II methanotrophs Methylocystis dominated in LCS. Methylobacter seemed tolerant to TCE and/or may play an important role in the TCE degradation in both soils, especially in WBS. These findings will

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Fig. 8. Cluster analysis of terminal-restriction fragments (T-RFs) of pmoA amplicons digested with Msp I on days 0 (original), 15, 30 and 54. BrayeCurtis similarity index was calculated using the abundance of T-RFs. The capital letters W and L in the label denote WBS and LCS, respectively; the number after the first dash represents the sampling time (days 0 (original), 15, 30 and 54); the number after the second dash represents the initial TCE concentration (ppmv).

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