Journal of Bioscience and Bioengineering VOL. 120 No. 6, 670e676, 2015 www.elsevier.com/locate/jbiosc
Inhibitory effects of sulfur compounds on methane oxidation by a methane-oxidizing consortium Eun-Hee Lee, Kyung-Eun Moon, Tae Gwan Kim, Sang-Don Lee, and Kyung-Suk Cho* Department of Environmental Science and Engineering, Ewha Womans University, 52 Ewhayeodae-gil, Seodaemun-gu, Seoul 120-750, Republic of Korea Received 26 December 2014; accepted 8 April 2015 Available online 2 July 2015
Kinetic and enzymatic inhibition experiments were performed to investigate the effects of methanethiol (MT) and hydrogen sulfide (H2S) on methane oxidation by a methane-oxidizing consortium. In the coexistence of MT and H2S, the oxidation of methane was delayed until MT and H2S were completely degraded. MT and H2S could be degraded, both with and without methane. The kinetic analysis revealed that the methane-oxidizing consortium showed a maximum methane oxidation rate (Vmax) of 3.7 mmol g-dry cell weight (DCW)L1 hL1 and a saturation constant (Km) of 184.1 mM. MT and H2S show competitive inhibition on methane oxidation, with inhibition values (Ki) of 1504.8 and 359.8 mM, respectively. MT was primary removed by particulate methane monooxygenases (pMMO) of the consortium, while H2S was degraded by the other microorganisms or enzymes in the consortium. DNA and mRNA transcript levels of the pmoA gene expressions were decreased to w106 and 103 pmoA gene copy number g-DCWL1 after MT and H2S degradation, respectively; however, both the amount of the DNA and mRNA transcript recovered their initial levels of w107 and 105 pmoA gene copy number g-DCWL1 after methane oxidation, respectively. The gene expression results indicate that the pmoA gene could be rapidly reproducible after methane oxidation. This study provides comprehensive information of kinetic interactions between methane and sulfur compounds. Ó 2015, The Society for Biotechnology, Japan. All rights reserved. [Key words: Methane; Particulate methane monooxygenases; Methanethiol; Hydrogen sulfide; Interaction]
Methane (CH4) is one of the most important global warming gases, with a global warming potential that is 20 times greater than that of carbon dioxide (CO2) (1). Methane is primarily generated by anaerobic degradation via methanogenesis and is produced in large amounts in landfills. Landfills have historically been the largest source of global methane emissions from the waste sector (1). Volatile sulfur compounds are emitted simultaneously with methane by the anaerobic digestion of sulfur-containing organic compounds (2,3). Methanethiol (CH3SH, MT) and hydrogen sulfide (H2S) are predominant landfill gases, strongly contributing to its malodor (2e4). For instance, Gendebien et al. reported a concentration of MT up to 87 mg m3 at British landfills (5). The United States Department of Energy summarized that H2S comprises 5.64 mmol Cu g-protein1 in the environment, as in landfills, forests, and wetlands (13e15). To date, most of the previous researches have focused on the cometabolic kinetics of volatile sulfur compounds by sMMO or by pure methanotrophic cultures (16e18). However, these studies have been unable to provide sufficient information of the kinetic interactions, since sMMO or pure cultures could not represent methanotrophic activity or community in environmental samples. In addition, earlier studies have only focused on the kinetic values, such as the maximum oxidation rates, saturation constants, or inhibition constants. This has led to a lack of information and limitations on our understanding of the underlying kinetic mechanisms. Accordingly, enzymatic and genetic approaches are required for a better understanding as well as kinetic analysis. In the present study, kinetic, enzymatic, and genetic approaches were employed to investigate the effects of MT and H2S on methane oxidation using a methane-oxidizing consortium. In order to
1389-1723/$ e see front matter Ó 2015, The Society for Biotechnology, Japan. All rights reserved. http://dx.doi.org/10.1016/j.jbiosc.2015.04.006
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examine the effects of the sulfur compounds, the methaneoxidizing consortium was obtained from landfill soil. The consortium was cultivated to express pMMO exclusively. Batch-scale experiments were employed to analyze the kinetic interactions between the sulfur compounds and the methane. Enzymatic assays were performed to evaluate the effects of MT and H2S on methane oxidation using the MMO of the methane-oxidizing consortium. The DNA level of pmoA gene (encoding the a subunit of pMMO) was measured using quantitative real-time PCR (q-PCR) and mRNA transcription level was analyzed using reverse transcriptase q-PCR The effects of MT and H2S on methane oxidation were comprehensively discussed at the kinetic, enzymatic, and genetic levels. MATERIALS AND METHODS Enrichment of a methane-oxidizing consortium In order to obtain a methane-oxidizing consortium, a landfill soil was sampled at Gongju-si, Chungcheongnam-do, South Korea. Two grams (wet weight) of the soil was added into a 600-ml serum bottle containing 18 ml of a nitrate mineral salt (NMS) medium with 0.03 mM of CuSO4. The bottle was sealed with a butyl rubber stopper and an aluminum cap, and methane gas was injected into the bottle at 5% (v/v) of the final concentration from a methane gas cylinder (99%, Seoul Special Gases, Seoul, Korea). The serum bottle was incubated at 30 C with an agitation of 200 rpm. The methane concentration was periodically monitored in the headspace, and the culture was transferred into the fresh NMS medium after the complete oxidation of methane. This process was repeated twenty times. The NMS medium consists of MgSO4$7H2O (1 g), CaCl2$2H2O (0.295 g), KNO3 (1 g), KH2PO4 (0.26 g), and Na2HPO4$2H2O (0.41 g per 1-L); plus trace elements of FeSO4$7H2O (500 mg), ZnSO4$7H2O (400 mg), MnCl2$4H2O (20 mg), H3BO4 (15 mg), CoCl2$6H2O (50 mg), NiCl2$6H2O (10 mg), and Na2MoO4$2H2O (250 mg) (19). The sampling site and physico-chemical properties of the soil were described in a previous report (20). The gram-dry cell weight (g-DCW) was measured to determine the cell concentration of the consortium. One ml of the culture was collected into the 1.5 ml tubes with a centrifugation at 13,000 rpm for 5 min, and dried at 80 C overnight. The g-DCW was determined from the residue in the tubes. All of the experiments were performed in triplicate. A bacterial community analysis of a methane-oxidizing consortium One milliliter of the culture was collected from the methane-oxidizing consortium. Genomic DNA was extracted in duplicate using a NucleoSpin Soil Kit (MachereyeNagel GmbH & Co. KG, Düren, Germany) with a modification as previously described (21). DNA was eluted in 100 mL of the elution buffer and stored at 20 C prior to use. DNA concentration was measured using a spectrophotometer (ASP-2680, ACTGene Inc., Piscataway, NJ, USA). In order to analyze the bacterial community of the methane-oxidizing consortium, the pyrosequencing assay was performed in duplicate. For PCR, a primer set of 340F and 805R was used in order to amplify the 16S rRNA gene. Six different composite primer sets were made, based on the primer set of 340F and 805R, for multiplex pyrosequencing (21). This procedure has been previously described in detail by Kim et al. (21). The purified DNA concentrations were quantified using a spectrophotometer. Equal amounts of the purified DNAs were combined in a single tube and run on a Genome Sequencer 454 FLX Titanium system (Roche Diagnostics Inc., Mannheim, Germany). For high-quality sequences, the primer sites of the sequences were trimmed and the low-quality and chimera sequences were removed. The primer sites and low-quality sequences (length < 400 nt, average quality score < 25, and with an ambiguity) were excluded using the ribosomal database project (RDP) pyrosequencing pipeline (22). Any possible chimeras were removed using Black Box Chimera Check software with the default settings (23). The RDP pyrosequencing pipeline was employed in order to analyze the pyrosequencing data, as previously described by Kim et al. (21). The sequences were identified at the species level, when the similarity was greater than 99%. The pyrosequencing reads obtained in this study were deposited into the DNA DataBank of Japan (DDBJ) Sequence Read Archive (http://trace.ddbj.nig.ac.jp/dra) under the accession no. DRA002267. Kinetic analysis for sulfur compounds effects on methane oxidation Kinetic analysis was performed to investigate the effects of sulfur compounds on methane oxidation by the methane-oxidizing consortium. Twenty milliliters of the enriched methane-oxidizing consortium was added into 600-ml serum bottles, and the bottles were sealed with butyl rubber stoppers and aluminum caps. Control tests were also performed without inoculation of the consortium. Methane, MT (99%, SigmaeAldrich, St. Louis, MO, USA), and H2S (99%, Seoul special gas, Seoul, Korea) gases were added to the bottles, for final concentrations of 27.2e204, 163.5e654, and 98.1e392.4 mmol L1, respectively. The liquid concentrations of methane, MT, and H2S were calculated using dimensionless Henry’s law constants of 29, 0.123, and 0.41, respectively (24,25). The serum bottles were incubated at 30 C with 200 rpm of an agitation. The
Unclassified Bradyrhizobiaceae
FIG. 1. A bacterial community structure of a methane-oxidizing consortium. The bacterial genera with assigned read numbers of 1% of the sequencing effort were excluded.
methane, MT, and H2S concentrations were periodically measured in the headspace using a gas-tight syringe. The methane, MT, and H2S degradation rates were calculated from the slopes of concentration plotted versus time. The maximum methane oxidation rate (Vmax) and saturation constant (Km) were determined from the LineweavereBurk equation below (26): 1 Km 1 1 ¼ þ V Vmax ½S Vmax
(1)
where V represents the methane oxidation rate and [S] represents the methane concentration. The inhibition constants (Ki) of MT and H2S on methane oxidation were calculated using the equation for competitive inhibition, given by Williams et al. (27): 1 Km ¼ V Vmax
½i 1 1 þ 1þ ½S Vmax Ki
(2)
where [i] represents the MT or H2S concentration. All of the experiments were performed in triplicate. Inhibitory effects of methanethiol and hydrogen sulfide on methane oxidation by MMO pMMO was exclusively induced for the methane oxidation of the consortium by cultivating the culture in the presence of Cu. The bottles were prepared, as described above, with the addition of allylthiourea (SigmaeAldrich). Allylthiourea has been widely used as an inhibitor for pMMO (28,29). Allylthiourea was supplemented into the bottles at 50 mM of the final concentration. Methane, MT, and H2S gases were added to the bottles, for final concentrations of 68, 327, and 196.2 mmol L1 of liquid, respectively. The serum bottles were incubated at 30 C with 200 rpm of an agitation. The concentrations of methane, MT, and H2S in the headspace were monitored every 4 or 5 h using a gas-tight syringe. All of the experiments were performed in triplicate. Quantitative analysis of the pmoA gene expression One milliliter of each culture was collected from the serum bottles of the kinetic experiments earlier at 48, 12, and 5 experimental hours, which marked the end of the degradation periods for CH4, MT, and H2S, respectively (Figs. 2 and 3). The samples were immediately frozen at 70 C prior to analysis. Genomic DNA was extracted from the collected samples in duplicate using a NucleoSpin Soil Kit (MachereyeNagel GmbH & Co. KG) as described above. The DNA was eluted in 100 mL of the elution buffer and stored at 20 C. RNA was extracted from the collected samples in duplicate. For RNA extraction, 1 ml of the culture was added to 2-mL microcentrifuge tubes containing 0.1-mm glass beads and 1-mm zirconia/silica beads (0.5 g each). This procedure was described in detail by Kim et al. (21). RNA pellets were suspended in 50 mL of 0.1% DEPC-treated water. RNA was immediately purified using a Qiagen RNeasy Mini kit (Qiagen Inc., Valencia, CA, USA) with a RNase-Free DNase set (Qiagen Inc.), as specified by the manufacturer. RNA extracts were quantified using a
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FIG. 2. The effect of methanethiol (MT) concentrations on methane oxidation by the methane-oxidizing consortium. Time profiles of methane (a) and MT concentration (b). Closed and open symbols represent methane and MT, respectively. Closed circles, methane alone (68 mM); triangles, methane (68 mM) þ MT (163.5 mM); squares, methane (68 mM) þ MT (327 mM); inverted triangles, methane (68 mM) þ MT (490.5 mM); diamonds, methane (68 mM) þ MT (654 mM); crosses, control (no inoculation).
spectrophotometer. All RNAs were stored at 70 C prior to use. DNA contamination was confirmed by PCR, using the 340F-805R primer set. The PCR mixture and operating conditions have previously been outlined by Kim et al. (21). The PCR results were checked using a 2% agarose gel electrophoresis. No positive results were observed from the RNA extracts (data not shown). The RNA was reverse-transcribed using a Qiagen Omniscript RT kit (Qiagen), according to the manufacturer’s recommendations. The 20-mL reaction mixture contained 1 RT Sensiscript buffer, dNTP at a concentration of 0.5 mM, a random hexamer at 10 mM, 10 U of RNase inhibitor (RNaseOUT, Invitrogen Inc., Grand Island, USA), 1 mL of RT-Sensiscript reverse transcriptase, and 10 mL of template RNA. q-PCR was employed to quantify the pmoA gene expression of the methanotrophs using the primer set of A189f and mb661r, which targets the pmoA gene (30). The pmoA gene of Methylobacter luteus (NCIMB11914) was used to establish a standard curve for the quantitative detection. The PCR mixture and operating conditions were previously described (20). DNA and cDNA were used for the q-PCR templates in order to determine the pmoA copy numbers of the target DNA and mRNA transcript. The q-PCR was performed in five replicates. Gas analysis Methane was measured using gas chromatography (GC, 6850N, Agilent Technologies Inc., Santa Clara, CA, USA) equipped with a flame ionization detector (FID) and a wax column (30 m 0.32 mm 0.25 mm, Supelco Inc., Bellefonte, PA, USA), as previously described (20). The oven, injector, and detector temperatures were 100 C, 230 C, and 230 C, respectively. MT and H2S were analyzed using a GC (7890A, Agilent Technologies Inc., Santa Clara, USA) equipped with a flame photometric detector (FPD) and a DB-1 column (30 m 0.3 mm, J&W Scientific, Folsom, CA, USA). The oven, injector, and detector temperatures were 130 C, 200 C, and 250 C, respectively. The detection limits were 10, 3, and 3 ppm for CH4, MT, and H2S, respectively. Statistical analysis A statistical analysis of variance (ANOVA) was performed to validate the q-PCR abundance of each group using the SPSS 20 for Windows (IBM Corp., Armonk, NY, USA) program. A regression analysis was employed to estimate
J. BIOSCI. BIOENG.,
FIG. 3. The effect of hydrogen sulfide (H2S) concentrations on methane oxidation by the methane-oxidizing consortium. Time profiles of methane (a) and H2S concentration (b). Closed and open symbols represent methane and H2S, respectively. Closed circles, methane alone (68 mM); triangles, methane (68 mM) þ H2S (98.1 mM); squares, methane (68 mM) þ H2S (196.2 mM); inverted triangles, methane (68 mM) þ H2S (294.3 mM); diamonds, methane (68 mM) þ H2S (392.4 mM); crosses, control (no inoculation).
the relationship between the methane concentration and the methane oxidation rate. The level of significance was p < 0.05 for all of the statistical analyses.
RESULTS A bacterial community structure of a methane-oxidizing consortium The bacterial community of the methane-oxidizing consortium was analyzed using pyrosequencing assay. As shown in Fig. 1, 12 bacterial genera (Steroidobacter, Sphingopyxis, Opitutus, Methylocystis, unclassified Methylocystaceae, Meiothermus, Hyphomicrobium, unclassified Gammaproteobacteria, unclassified Chitinophagaceae, unclassified Bradyrhizobiaceae, Bosea, and unclassified Beijerinckiaceae) were detected in the consortium. The genus Methylocystis dominated the bacterial community in the consortium, followed by unclassified Chitinophagaceae, unclassified Gammaproteobacteria, Meiothermus, and Sphingopyxis. Methylocystis comprised 51 0.2% of the bacterial community in the methane-oxidizing consortium. Kinetic analysis for methanethiol and hydrogen sulfide effects on methane oxidation Fig. 2 shows the methane oxidation and MT degradation in the consortium. The consortium was capable of oxidizing methane without a lag period at a methane concentration of 68 mM in the absence of MT. However, methane oxidation was delayed in the presence of MT, and no methane oxidation was observed during an experimental period
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FIG. 4. The effect of methanethiol (MT) and hydrogen sulfide (H2S) on the methane oxidation rates by a methane-oxidizing consortium at different methane concentrations.
at the MT concentrations above 490.5 mM (Fig. 2a). The methane oxidation rates decreased with increasing MT concentrations, from 1134.0 9.6 to 773.2 29.2 mmol g-dry cell weight (DCW)1 h1 at the MT concentrations of 0e327 mM. On the other hand, MT was degraded immediately by the methaneoxidizing consortium, and the degradation occurred even at a high concentration of 654 mM of MT without inhibition (Fig. 2b). The MT degradation rates showed 13.6e4.8 mmol g-DCW1 h1 at 327e654 mM of MT, which decreased as MT concentrations increased. The addition of H2S inhibited methane oxidation of the methane-oxidizing consortium (Fig. 3). The period for methane oxidation increased with increasing H2S concentrations, for instance, it took 4 days to oxidize approximately 90% of the methane in the presence of 392.4 mM H2S (Fig. 3a). The methane oxidation rates decreased from 1085.7 8.6 to 734.2 18.2 mmol gDCW1 h1 at the H2S concentrations of 0e196.2 mM. The consortium degraded H2S at concentrations of 98.1e392.4 mM without any delays. The H2S degradation rates ranged from 17.5 to 81.0 mmol g-DCW1 h1 at 98.1e392.4 mM of H2S, which were greater than the rates of MT degradation (Figs. 2b and 3b). H2S was completely degraded within 5 h, even at a high concentration of 392.4 mM (Fig. 3b). Fig. 4 shows the effect of MT and H2S on methane oxidation at different concentrations of methane. The methane oxidation rates of the consortium were measured at different methane concentrations, ranging from 27.2 to 204 mM. Then, they were compared with the methane oxidation rates which were measured in the presence of MT (327 mM) or H2S (196.2 mM). MT addition lead to a decrease of 13e31% for the methane oxidation, compared to the methane oxidation rates at 27.2e136 mM of methane alone. H2S addition caused a 31e38% decrease in the methane oxidation rates, compared to the rates at 27.2e136 mM of methane alone. However, no significant decrease of methane oxidation rates was observed at 204 mM of methane under the coexistence of MT or H2S. LineweavereBurk plots (Eq. 1) were constructed to estimate the Vmax, Km, and Ki values, based on MichaeliseMenten kinetics (data not shown). The methane-oxidizing consortium showed 3.7 mmol g-DCW1 h1 of Vmax and 184.1 mM of Km. The Ki values were determined from Eq. 2 for competitive inhibition (27), since it showed similar Vmax values, irrespective of MT and H2S addition. Ki parameters were calculated using Eq. 2 with the aforementioned
Hydrogen sulfide concentration in headspace (ppm)
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Time (h) FIG. 5. Effects of allylthiourea on the degradation of methanethiol (MT) and hydrogen sulfide (H2S) by a methane-oxidizing consortium. Closed and open symbols represent MT or H2S degradation in the absence and presence of allylthiourea, respectively. (a) Time profile of MT concentration. Circles, MT alone (327 mM); squares, MT (327 mM) þ methane (68 mM); closed inverted triangle, control (no inoculation). (b) Time profile of H2S concentration. Circles, H2S alone (196.2 mM); squares, H2S (196.2 mM) þ methane (68 mM); closed inverted triangle, control (no inoculation).
values of Vmax and Km. The Ki values of MT and H2S were 1504.8 and 359.8 mM, respectively. Inhibitory effects of methanethiol and hydrogen sulfide on methane oxidation by MMO The MMO assay was performed to investigate the effects of MT and H2S on methane oxidation by the methane-oxidizing consortium. Particularly, pMMO was targeted for the assay, since the consortium was induced to express pMMO exclusively by adding Cu ions into the media during cultivation. Allylthiourea is a well-known inhibitor on the synthesis of pMMO (28,29). Yu et al. (29) reported that allylthiourea addition reduced the amount of intracellular copper by preventing entry of copper into the cells. This presumably leads to the decrease of the methanotrophic pMMO expression. No methane oxidation occurred in the presence of allylthiourea (data not shown). This indicated that methane was oxidized exclusively by the pMMO of the methane-oxidizing consortium. The MT was completely degraded within 12 h in the absence of
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FIG. 6. Effects of methanethiol (MT) and hydrogen sulfide (H2S) on the pmoA gene expression in the methane-oxidizing consortium. The pmoA gene levels were analyzed using cultures that were sampled from the serum bottles of the kinetic experiments (Figs. 2 and 3). In the coexistence of methane with MT and H2S, the culture was collected after 5, 12, 48 h of incubation time (shown in Figs. 2 and 3), which are ends of degradation periods for H2S, MT, and CH4, respectively. The initial concentrations of CH4, MT, and H2S were 68, 327, and 196.2 mM respectively. (a) Abundance of pmoA gene at DNA level. (b) Abundance of pmoA gene at mRNA transcript level.
allylthiourea (Fig. 5a). However, MT degradation was retarded in the presence of allylthiourea and only 41e44% of the MT was removed for 12 h. H2S was degraded by the consortium, irrespective of allylthiourea addition, showing a slight increase in the presence of allylthiourea (Fig. 5b). H2S was degraded 99.6 0.5% in the presence of allylthiourea within 6.5 h, while it was 89.0 1.9% in the absence of allylthiourea. Quantification of pmoA gene expression in the methaneoxidizing consortium q-PCR was performed to quantify the
pmoA gene copy numbers in the methane-oxidizing consortium using DNA and RNA extracts from the kinetic experiments. The initial DNA and mRNA transcript levels of the pmoA gene were 9.9 1.2 106 and 1.6 0.7 105 pmoA gene copy number gDCW1, respectively (Fig. 6). The DNA levels of the pmoA gene decreased to 2.6 0.2 106 and 3.5 0.2 106 pmoA gene copy number g-DCW1 after its degradation for MT or H2S alone, respectively, whereas it was slightly increased to 1.6 0.1 107 pmoA gene copy number g-DCW1 after methane oxidation for methane alone (Fig. 6a). In the coexistence of methane with MT and H2S, the DNA levels of the pmoA gene decreased to 5.4 0.32 106 and 2.1 1.0 106 pmoA gene copy number gDCW1 after MT and H2S degradation (at the point of 12 and 5 h in incubation time), respectively, methane oxidation recovered to the initial DNA level of w107 pmoA gene copy number g-DCW1 (at the point of 48 h in incubation time). No statistical differences were observed among the DNA levels of the pmoA gene as compared to the initial value. In contrast, the amount of mRNA transcript for the pmoA gene expression changed significantly between the samples (Fig. 6b). For example, in the coexistence of methane with MT and H2S, the mRNA transcript levels of the pmoA gene decreased greatly to 3.8 4.2 103 and 6.6 1.2 103 pmoA gene copy number g-DCW1 after MT and H2S degradation (at the point of 12 and 5 h in incubation time), respectively, compared to the initial level, and they were subsequently recovered to 2.2e6.6 0.7 105 pmoA gene copy number g-DCW1 after methane oxidation (at the point of 48 h in incubation time). The decrease and recovery of the mRNA transcript levels of the pmoA gene were statistically proven using ANOVA analysis.
DISCUSSION Kinetic and enzymatic approaches were performed to investigate the effects of MT and H2S on methane oxidation by the methane-oxidizing consortium. The consortium obtained in this study oxidized methane successfully without a lag period, since the methanotrophs were actively cultured by continuously enriching the consortium with methane as the sole carbon and energy source (Figs. 2 and 3). Methanotrophs are classified into two major groups of type I and type II, depending on the DNA content, intracellular membrane arrangement, carbon assimilation pathway, and PLFA composition (31). In this study, the type II methanotroph Methylocystis dominated the bacterial community, comprising 51 0.2% of the bacterial community in the methane-oxidizing consortium (Fig. 1). Generally, the methane concentration can affect methanotrophic community compositions (32,33). The consortium was continuously exposed to a relatively high methane concentration of 68 mM during enrichment. This condition supplied a favorable environment to type II methanotrophs and this could lead to the dominance of Methylocystis in the bacterial community. Our observations support previous studies, which reported that type II methanotrophs preferred methane rich environments and outcompeted type I methanotrophs in high methane concentrations (32,34). Methylocystis is a well-known methane-oxidizing bacterium (8), indicating that this genus plays a primary role in methane oxidation. The MMO assay revealed that the methane was oxidized exclusively by pMMO (data not shown). This is not surprising, as the consortium was continuously cultured in an NMS medium supplemented with Cu ions. Generally, Cu regulates MMO expression in methanotrophs and the pMMO is expressed at high concentrations of Cu >0.8 mmol of Cu per g-DCW (8). The consortium showed 3.7 mmol g-DCW1 h1 of Vmax and 184.1 mM of Km. The Vmax and Km values are greater than or comparable to the levels observed in previous studies. For instance,
VOL. 120, 2015 Choi et al. (35) reported the Vmax and Km values for methane oxidation by a mixed culture of methanotrophs were 326.8 mmol of CH4 g-DCW1 h1 and 143.8 mM, respectively. Lee et al. (17) calculated Vmax and Km values for methane oxidation by Methylocystis sp. M6 as 4.93 mmol of CH4 g-DCW1 h1 and 23 mM, respectively. Kinetic analysis revealed that MT and H2S had inhibitory effects on methane oxidation by the consortium (Figs. 2 and 3). Methane oxidation was delayed in the presence of MT or H2S, and their methane oxidation rates decreased to 424.4e773.2 and 330.0e484.3 mmol g-DCW1 h1, respectively, corresponding to a 13e31 and 30e50% decrease of the oxidation rates at 27.2e68 mM of methane alone (Figs. 2e4). This is consistent with earlier studies, which reported that the methane oxidation of Methylocysitis sp. M6 was inhibited by the presence of MT and H2S (17). Börjesson (36) studied the inhibition of methane oxidation by volatile sulfur compounds in landfill cover soils, describing MT’s inhibitory effects on methane oxidation rates. Jäckel et al. (37) found that the addition of MT resulted in an inhibition of methane oxidation in deciduous forest soil. Cáceres et al. (16) reported that H2S over 0.05% (v/v) decreased the methane oxidation rate of Methylocystis sp. The expression of pMMO in the consortium was quantified using a q-PCR with DNA and RNA extracts from the kinetic experiments. The results of the pmoA gene expression also support the kinetic analysis (Fig. 6). In the coexistence of methane with MT and H2S, the DNA and mRNA transcript levels for the pmoA gene expression were decreased to 2.1e5.4 106 and 3.8e6.6 103 pmoA gene copy number g-DCW1 after MT (at the point of 12 h in incubation time) and H2S degradation (at the point of 5 h in incubation time), respectively, as compared to their initial levels. However, both the amount of DNA and mRNA transcript for the pmoA gene were recovered to the initial levels of w107 and 105 pmoA gene copy number g-DCW1 after methane oxidation (at the point of 48 h in incubation time), respectively. Interestingly, the changes in the pmoA gene expression differed between the results of the DNA- and RNA-based analyses. Although the DNA levels of the pmoA gene were decreased after MT and H2S degradation, the differences were observed on the same order of 106 level, whereas the mRNA transcripts showed a 100-fold decrease. These results were confirmed by a statistical analysis. No difference was observed between the DNA levels of the pmoA gene; however, the mRNA transcripts levels varied significantly. The analysis of DNA-based gene expression can be used for evaluation of microbial activities and microbial ecology, however that provides relatively little information since DNA is known to be stable even in dead cells (38). On the other hand, the analysis of mRNA-based gene expression can provide relatively more information on the microbial activities and ecological significance as it has short half-life (wminutes) which is primary regulated at the level of transcription (39e41). For instance, Dumont et al. (42) found that the C13 labelling of the mRNA for the pmoA gene was quicker than DNA, following the incubation of lake sediment with 13CH4, demonstrating that mRNA-stable isotope probing (SIP) is more sensitive than DNA-SIP. Han and Semrau (43) examined the quantification of gene expression in methanotrophs using a competitive reverse transcription-polymerase chain reaction and reported that the amount of mRNA transcript for the pmoA gene correlated well with the whole-cell pMMO activity. In this study, the expression of pmoA gene was quantified at the point of 5, 12, and 48 h in incubation time of the kinetic experiments, this presumably only induced the changes of the mRNA-based pmoA gene levels because of short time of incubation. Based on the previous reports and our results, it is possible to conclude that MT and H2S dramatically inhibit the pMMO activity of the consortium since mRNA levels were decreased due to MT and H2S addition, resulting in a decrease of methane oxidation by repressing pmoA gene expression. A comparison between the results of DNA- and RNA-
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based pmoA gene levels demonstrated that the DNA results alone can lead to an underestimation of the active gene expression since it is probably incapable of sensitively reflecting changes of pMMO activity. The LineweavereBurk plots revealed that MT and H2S follow a typical competitive inhibition model, since they had the same yintercepts as those of methane alone (data not shown). As shown in Fig. 4, the methane oxidation rates were similar at 204 mM of methane, regardless of whether MT or H2S were added. This means that if the amount of methane is very high, the effect of the inhibitors (MT and H2S) becomes less noticeable. The inhibition constants of MT and H2S were 1504.8 and 359.8 mM, respectively, determined using Eq. 2 for competitive inhibition. These values indicate that H2S has greater inhibitory effects on methane oxidation than MT. This is consistent with an earlier study for the inhibitory effects of MT on methane oxidation, which followed a competitive inhibition model with an inhibition constant of 40.6 18.5 nmol or 1.95 mg MT g-dry soil1 (36). Although MT and H2S inhibited methane oxidation by the methane-oxidizing consortium, methane oxidation occurred in the presence of MT (327 mM) and H2S (196.2 mM). This suggests that MT and H2S are possibly classified as reversible inhibitors. The results of the mRNA transcript levels for the pmoA gene expression also support this observation, which were recovered to the level of 105 pmoA gene copy number g-DCW1 after methane oxidation (Fig. 6). The methane-oxidizing consortium was capable of degrading MT and H2S, and MT and H2S degradation occurred without a lag period (Figs. 2, 3, 5, and 6). MT degradation rapidly decreased in the presence of allylthiourea, indicating that most of the MT was degraded by pMMO (Fig. 5a). This suggests that methanotrophs play an important role in MT degradation. In addition, MT was slightly degraded in the presence of allylthiourea, compared to the abiotic experiment (control), suggesting that other microorganisms or enzymes were marginally involved in the MT degradation as well. Conversely, H2S degradation occurred, irrespective of allylthiourea addition, indicating that pMMO was not associated with the H2S degradation (Fig. 5b). Interestingly, H2S was completely degraded by the consortium, suggesting that other microorganisms or enzymes play a primary role in the H2S degradation. Thus, we can conclude that methanotrophs are capable of cometabolizing MT, but not H2S, while other microorganisms or enzymes are primary involved in the H2S degradation. Börjesson (36) observed that MT was rapidly consumed, with a rate of 16.03 3.6 mmol MT g-dry soil1 h1, in landfill cover soils. Zhang et al. (44) reported that Hyphomicrobium sp. is capable of removing MT and H2S. Lee et al. (45) found that Sphingopyxis sp. degrades MT and H2S. These studies indicate that the genera Hyphomicrobium and Sphingopyxis have potential for MT and H2S degradation in the consortium (Fig. 1). The objective of this study was to investigate the kinetic and enzymatic inhibitory effects of MT and H2S on methane oxidation of the consortium. MT and H2S showed competitive inhibition on methane oxidation. MT was primary degraded by the cometabolism of pMMO, whereas H2S was degraded by other microorganisms or enzymes in the consortium. Our findings demonstrated that pMMO is considerable for the cometabolism of MT, despite of its narrow range of substrate specificity than sMMO, since the pMMO was easily reproducible (as shown in the analysis of pmoA gene expression). On the basis of the results, we conclude that this study provides valuable information for methane mitigation and its kinetics and interactions with sulfur compounds. ACKNOWLEDGMENTS This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF),
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funded by the Ministry of Science, ICT and Future Planning (NRF2012R1A2A2A03046724), and by the National Research Foundation of Korea (NRF 2009-83527). E. H. Lee was financially supported by National Research Foundation of Korea (NRF-2014R1A1A3051952). References 1. Solomon, S.: Climate change 2007-the physical science basis: Working group I contribution to the fourth assessment report of the IPCC. Cambridge University Press (2007). 2. Smet, E. and Van Langenhove, H.: Abatement of volatile organic sulfur compounds in odorous emissions from the bio-industry, Biodegradation, 9, 273e284 (1998). 3. Themelis, N. J. and Ulloa, P. A.: Methane generation in landfills, Renew. Energy, 32, 1243e1257 (2007). 4. Saral, A., Demir, S., and Yildiz, S.: Assessment of odorous VOCs released from a main MSW landfill site in Istanbul-Turkey via a modelling approach, J. Hazard. Mater., 168, 338e345 (2009). 5. Gendebien, A., Pauwels, M., Constant, M., Ledrut-Damanet, M.-J., Nyns, E.-J., Willumsen, H.-C., Butson, J., Fabry, R., and Ferrero, G.-L.: Landfill gas. From environment to energy. Commission of the European Communities, Luxembourg (1992). 6. U.S. Environmental Protection Agency: Municipal solid waste landfills, section 2.4, in: AP-42, 5th ed. Compilation of air pollutant emissions factors. Volume I: Stationary point and area sources. U.S. Environmental Protection Agency, Research Triangle Park, NC (1995). 7. Lassey, K. R., Etheridge, D. M., Lowe, D. C., Smith, A. M., and Ferretti, D. F.: Centennial evolution of the atmospheric methane budget: what do the carbon isotopes tell us? Atmos. Chem. Phys., 7, 2119e2139 (2007). 8. Hanson, R. S. and Hanson, T. E.: Methanotrophic bacteria, Microbiol. Rev., 60, 439e471 (1996). 9. Vorobev, A. V., Baani, M., Doronina, N. V., Brady, A. L., Liesack, W., Dunfield, P. F., and Dedysh, S. N.: Methyloferula stellata gen. nov., sp. nov., an acidophilic, obligately methanotrophic bacterium that possesses only a soluble methane monooxygenase, Int. J. Syst. Evol. Microbiol., 61, 2456e2463 (2011). 10. McDonald, I. R., Bodrossy, L., Chen, Y., and Murrell, J. C.: Molecular ecology techniques for the study of aerobic methanotrophs, Appl. Environ. Microbiol., 74, 1305e1315 (2008). 11. Murrell, J. C., McDonald, I. R., and Gilbert, B.: Regulation of expression of methane monooxygenases by copper ions, Trends Microbiol., 8, 221e225 (2000). 12. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C.: Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2, Mol. Microbiol., 58, 682e692 (2005). 13. Hakemian, A. S. and Rosenzweig, A. C.: The biochemistry of methane oxidation,, Annu. Rev. Biochem., 76, 223e241 (2007). 14. Semrau, J. D., DiSpirito, A. A., and Yoon, S.: Methanotrophs and copper, FEMS Microbiol. Rev., 34, 496e531 (2010). 15. Morton, J. D., Hayes, K. F., and Semrau, J. D.: Effect of copper speciation on whole-cell soluble methane monooxygenase activity in Methylosinus trichosporium OB3b, Appl. Environ. Microbiol., 66, 1730e1733 (2000). 16. Caceres, M., Gentina, J. C., and Aroca, G.: Oxidation of methane by Methylomicrobium album and Methylocystis sp. in the presence of H2S and NH3, Biotechnol. Lett., 36, 69e74 (2014). 17. Lee, E. H., Yi, T., Moon, K. E., Park, H., Ryu, H. W., and Cho, K. S.: Characterization of methane oxidation by a methanotroph isolated from a landfill cover soil, South Korea, J. Microbiol. Biotechnol., 21, 753e756 (2011). 18. Colby, J., Stirling, D. I., and Dalton, H.: The soluble methane mono-oxygenase of Methylococcus capsulatus (Bath). Its ability to oxygenate n-alkanes, n-alkenes, ethers, and alicyclic, aromatic and heterocyclic compounds, Biochem. J., 165, 395e402 (1977). 19. Whittenbury, R. and Dalton, H.: The methylotrophic bacteria, pp. 894e902, in: Starr, M., Stolp, H., Trüper, H., Balows, A., and Schlegel, H. (Eds.), The prokaryotes. Springer Berlin Heidelberg (1981). 20. Lee, E. H., Park, H., and Cho, K. S.: Characterization of methane, benzene and toluene-oxidizing consortia enriched from landfill and riparian wetland soils, J. Hazard. Mater., 184, 313e320 (2010). 21. Kim, T. G., Moon, K. E., Yun, J., and Cho, K. S.: Comparison of RNA- and DNAbased bacterial communities in a lab-scale methane-degrading biocover, Appl. Microbiol. Biotechnol., 97, 3171e3181 (2013).
J. BIOSCI. BIOENG., 22. Cole, J. R., Wang, Q., Cardenas, E., Fish, J., Chai, B., Farris, R. J., Kulam-SyedMohideen, A. S., McGarrell, D. M., Marsh, T., Garrity, G. M., and Tiedje, J. M.: The ribosomal database project: improved alignments and new tools for rRNA analysis, Nucleic Acids Res., 37, D141eD145 (2009). 23. Gontcharova, V., Youn, E., Wolcott, R. D., Hollister, E. B., Gentry, T. J., and Dowd, S. E.: Black box chimera check (B2C2): a Windows-based software for batch depletion of chimeras from bacterial 16S rRNA gene datasets, Open Microbiol. J., 4, 47e52 (2010). 24. Sander, R.: Compilation of Henry’s law constants for inorganic and organic species of potential importance in environmental chemistry (version 3), http:// www.henrys-law.org/henry-3.0.pdf. Max-Planck Institute, Mainz (1999). 25. Staudinger, J. and Roberts, P. V.: A critical compilation of Henry’s law constant temperature dependence relations for organic compounds in dilute aqueous solutions, Chemosphere, 44, 561e576 (2001). 26. Lineweaver, H. and Burk, D.: The determination of enzyme dissociation constants, J. Am. Chem. Soc., 56, 658e666 (1934). 27. Williams, V. R., Mattice, W. L., and Williams, H. B.: Kinetics, basic physical chemistry for the life sciences. WH Freeman and Company, San Fransisco, USA (1978). 28. Hubley, J., Thomson, A., and Wilkinson, J.: Specific inhibitors of methane oxidation in Methylosinus trichosporium, Arch. Microbiol., 102, 199e202 (1975). 29. Yu, Y. H., Ramsay, J., and Ramsay, B.: Use of allylthiourea to produce soluble methane monooxygenase in the presence of copper, Appl. Microbiol. Biotechnol., 82, 333e339 (2009). 30. Kolb, S., Knief, C., Stubner, S., and Conrad, R.: Quantitative detection of methanotrophs in soil by novel pmoA-targeted real-time PCR assays, Appl. Environ. Microbiol., 69, 2423e2429 (2003). 31. Bull, I. D., Parekh, N. R., Hall, G. H., Ineson, P., and Evershed, R. P.: Detection and classification of atmospheric methane oxidizing bacteria in soil, Nature, 405, 175e178 (2000). 32. Amaral, J. A. and Knowles, R.: Growth of methanotrophs in methane and oxygen counter gradients, FEMS Microbiol. Lett., 126, 215e220 (1995). 33. Graham, D., Chaudhary, J., Hanson, R., and Arnold, R.: Factors affecting competition between type I and type II methanotrophs in two-organism, continuous-flow reactors, Microb. Ecol., 25, 1e17 (1993). 34. Henckel, T., Roslev, P., and Conrad, R.: Effects of O2 and CH4 on presence and activity of the indigenous methanotrophic community in rice field soil, Environ. Microbiol., 2, 666e679 (2000). 35. Choi, S. A., Lee, E. H., and Cho, K. S.: Effect of trichloroethylene and tetrachloroethylene on methane oxidation and community structure of methanotrophic consortium, J. Environ. Sci. Health A, 48, 1723e1731 (2013). 36. Börjesson, G.: Inhibition of methane oxidation by volatile sulfur compounds (CH3SH and CS2) in landfill cover soils, Waste Manag. Res., 19, 314e319 (2001). 37. Jäckel, U., Schnell, S., and Conrad, R.: Microbial ethylene production and inhibition of methanotrophic activity in a deciduous forest soil, Soil Biol. Biochem., 36, 835e840 (2004). 38. Lindahl, T.: Instability and decay of the primary structure of DNA, Nature, 362, 709e715 (1993). 39. Kusher, S. R.: mRNA decay, pp. 849e860, in: Neidhardt, F. C., Curtiss, R., Ingraham, J. L., Lin, E. C. C., Low, K. B., and Magasanik, B. (Eds.), Escherichia coli and Salmonella: Cellular and molecular biology. American Society for Microbiology, Washington, D.C (1996). 40. Sheridan, G. E. C., Masters, C. I., Shallcross, J. A., and Mackey, B. M.: Detection of mRNA by reverse transcription PCR as an indicator of viability in Escherichia coli cells, Appl. Environ. Microbiol., 64, 1313e1318 (1998). 41. Spring, S., Schulze, R., Overmann, J., and Schleifer, K.: Identification and characterization of ecologically significant prokaryotes in the sediment of freshwater lakes: molecular and cultivation studies, FEMS Microbiol. Rev., 24, 573e590 (2000). 42. Dumont, M. G., Pommerenke, B., Casper, P., and Conrad, R.: DNA-, rRNA- and mRNA-based stable isotope probing of aerobic methanotrophs in lake sediment, Environ. Microbiol., 13, 1153e1167 (2011). 43. Han, J. I. and Semrau, J. D.: Quantification of gene expression in methanotrophs by competitive reverse transcription-polymerase chain reaction, Environ. Microbiol., 6, 388e399 (2004). 44. Zhang, L., Hirai, M., and Shoda, M.: Removal characteristics of dimethyl sulfide, methanethiol and hydrogen sulfide by Hyphomicrobium sp. 155 isolated from Peat Biofilter, J. Ferment. Bioeng., 72, 392e396 (1991). 45. Lee, J. H., Kim, T. G., and Cho, K. S.: Isolation and characterization of a facultative methanotroph degrading malodor-causing volatile sulfur compounds, J. Hazard. Mater., 235e236, 224e229 (2012).
Inhibitory effects of sulfur compounds on methane oxidation by a methane-oxidizing consortium Eun-Hee Lee, Kyung-Eun Moon, Tae Gwan Kim, Sang-Don Lee, and Kyung-Suk Cho* Department of Environmental Science and Engineering, Ewha Womans University, 52 Ewhayeodae-gil, Seodaemun-gu, Seoul 120-750, Republic of Korea Received 26 December 2014; accepted 8 April 2015 Available online 2 July 2015
Kinetic and enzymatic inhibition experiments were performed to investigate the effects of methanethiol (MT) and hydrogen sulfide (H2S) on methane oxidation by a methane-oxidizing consortium. In the coexistence of MT and H2S, the oxidation of methane was delayed until MT and H2S were completely degraded. MT and H2S could be degraded, both with and without methane. The kinetic analysis revealed that the methane-oxidizing consortium showed a maximum methane oxidation rate (Vmax) of 3.7 mmol g-dry cell weight (DCW)L1 hL1 and a saturation constant (Km) of 184.1 mM. MT and H2S show competitive inhibition on methane oxidation, with inhibition values (Ki) of 1504.8 and 359.8 mM, respectively. MT was primary removed by particulate methane monooxygenases (pMMO) of the consortium, while H2S was degraded by the other microorganisms or enzymes in the consortium. DNA and mRNA transcript levels of the pmoA gene expressions were decreased to w106 and 103 pmoA gene copy number g-DCWL1 after MT and H2S degradation, respectively; however, both the amount of the DNA and mRNA transcript recovered their initial levels of w107 and 105 pmoA gene copy number g-DCWL1 after methane oxidation, respectively. The gene expression results indicate that the pmoA gene could be rapidly reproducible after methane oxidation. This study provides comprehensive information of kinetic interactions between methane and sulfur compounds. Ó 2015, The Society for Biotechnology, Japan. All rights reserved. [Key words: Methane; Particulate methane monooxygenases; Methanethiol; Hydrogen sulfide; Interaction]
Methane (CH4) is one of the most important global warming gases, with a global warming potential that is 20 times greater than that of carbon dioxide (CO2) (1). Methane is primarily generated by anaerobic degradation via methanogenesis and is produced in large amounts in landfills. Landfills have historically been the largest source of global methane emissions from the waste sector (1). Volatile sulfur compounds are emitted simultaneously with methane by the anaerobic digestion of sulfur-containing organic compounds (2,3). Methanethiol (CH3SH, MT) and hydrogen sulfide (H2S) are predominant landfill gases, strongly contributing to its malodor (2e4). For instance, Gendebien et al. reported a concentration of MT up to 87 mg m3 at British landfills (5). The United States Department of Energy summarized that H2S comprises 5.64 mmol Cu g-protein1 in the environment, as in landfills, forests, and wetlands (13e15). To date, most of the previous researches have focused on the cometabolic kinetics of volatile sulfur compounds by sMMO or by pure methanotrophic cultures (16e18). However, these studies have been unable to provide sufficient information of the kinetic interactions, since sMMO or pure cultures could not represent methanotrophic activity or community in environmental samples. In addition, earlier studies have only focused on the kinetic values, such as the maximum oxidation rates, saturation constants, or inhibition constants. This has led to a lack of information and limitations on our understanding of the underlying kinetic mechanisms. Accordingly, enzymatic and genetic approaches are required for a better understanding as well as kinetic analysis. In the present study, kinetic, enzymatic, and genetic approaches were employed to investigate the effects of MT and H2S on methane oxidation using a methane-oxidizing consortium. In order to
1389-1723/$ e see front matter Ó 2015, The Society for Biotechnology, Japan. All rights reserved. http://dx.doi.org/10.1016/j.jbiosc.2015.04.006
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examine the effects of the sulfur compounds, the methaneoxidizing consortium was obtained from landfill soil. The consortium was cultivated to express pMMO exclusively. Batch-scale experiments were employed to analyze the kinetic interactions between the sulfur compounds and the methane. Enzymatic assays were performed to evaluate the effects of MT and H2S on methane oxidation using the MMO of the methane-oxidizing consortium. The DNA level of pmoA gene (encoding the a subunit of pMMO) was measured using quantitative real-time PCR (q-PCR) and mRNA transcription level was analyzed using reverse transcriptase q-PCR The effects of MT and H2S on methane oxidation were comprehensively discussed at the kinetic, enzymatic, and genetic levels. MATERIALS AND METHODS Enrichment of a methane-oxidizing consortium In order to obtain a methane-oxidizing consortium, a landfill soil was sampled at Gongju-si, Chungcheongnam-do, South Korea. Two grams (wet weight) of the soil was added into a 600-ml serum bottle containing 18 ml of a nitrate mineral salt (NMS) medium with 0.03 mM of CuSO4. The bottle was sealed with a butyl rubber stopper and an aluminum cap, and methane gas was injected into the bottle at 5% (v/v) of the final concentration from a methane gas cylinder (99%, Seoul Special Gases, Seoul, Korea). The serum bottle was incubated at 30 C with an agitation of 200 rpm. The methane concentration was periodically monitored in the headspace, and the culture was transferred into the fresh NMS medium after the complete oxidation of methane. This process was repeated twenty times. The NMS medium consists of MgSO4$7H2O (1 g), CaCl2$2H2O (0.295 g), KNO3 (1 g), KH2PO4 (0.26 g), and Na2HPO4$2H2O (0.41 g per 1-L); plus trace elements of FeSO4$7H2O (500 mg), ZnSO4$7H2O (400 mg), MnCl2$4H2O (20 mg), H3BO4 (15 mg), CoCl2$6H2O (50 mg), NiCl2$6H2O (10 mg), and Na2MoO4$2H2O (250 mg) (19). The sampling site and physico-chemical properties of the soil were described in a previous report (20). The gram-dry cell weight (g-DCW) was measured to determine the cell concentration of the consortium. One ml of the culture was collected into the 1.5 ml tubes with a centrifugation at 13,000 rpm for 5 min, and dried at 80 C overnight. The g-DCW was determined from the residue in the tubes. All of the experiments were performed in triplicate. A bacterial community analysis of a methane-oxidizing consortium One milliliter of the culture was collected from the methane-oxidizing consortium. Genomic DNA was extracted in duplicate using a NucleoSpin Soil Kit (MachereyeNagel GmbH & Co. KG, Düren, Germany) with a modification as previously described (21). DNA was eluted in 100 mL of the elution buffer and stored at 20 C prior to use. DNA concentration was measured using a spectrophotometer (ASP-2680, ACTGene Inc., Piscataway, NJ, USA). In order to analyze the bacterial community of the methane-oxidizing consortium, the pyrosequencing assay was performed in duplicate. For PCR, a primer set of 340F and 805R was used in order to amplify the 16S rRNA gene. Six different composite primer sets were made, based on the primer set of 340F and 805R, for multiplex pyrosequencing (21). This procedure has been previously described in detail by Kim et al. (21). The purified DNA concentrations were quantified using a spectrophotometer. Equal amounts of the purified DNAs were combined in a single tube and run on a Genome Sequencer 454 FLX Titanium system (Roche Diagnostics Inc., Mannheim, Germany). For high-quality sequences, the primer sites of the sequences were trimmed and the low-quality and chimera sequences were removed. The primer sites and low-quality sequences (length < 400 nt, average quality score < 25, and with an ambiguity) were excluded using the ribosomal database project (RDP) pyrosequencing pipeline (22). Any possible chimeras were removed using Black Box Chimera Check software with the default settings (23). The RDP pyrosequencing pipeline was employed in order to analyze the pyrosequencing data, as previously described by Kim et al. (21). The sequences were identified at the species level, when the similarity was greater than 99%. The pyrosequencing reads obtained in this study were deposited into the DNA DataBank of Japan (DDBJ) Sequence Read Archive (http://trace.ddbj.nig.ac.jp/dra) under the accession no. DRA002267. Kinetic analysis for sulfur compounds effects on methane oxidation Kinetic analysis was performed to investigate the effects of sulfur compounds on methane oxidation by the methane-oxidizing consortium. Twenty milliliters of the enriched methane-oxidizing consortium was added into 600-ml serum bottles, and the bottles were sealed with butyl rubber stoppers and aluminum caps. Control tests were also performed without inoculation of the consortium. Methane, MT (99%, SigmaeAldrich, St. Louis, MO, USA), and H2S (99%, Seoul special gas, Seoul, Korea) gases were added to the bottles, for final concentrations of 27.2e204, 163.5e654, and 98.1e392.4 mmol L1, respectively. The liquid concentrations of methane, MT, and H2S were calculated using dimensionless Henry’s law constants of 29, 0.123, and 0.41, respectively (24,25). The serum bottles were incubated at 30 C with 200 rpm of an agitation. The
Unclassified Bradyrhizobiaceae
FIG. 1. A bacterial community structure of a methane-oxidizing consortium. The bacterial genera with assigned read numbers of 1% of the sequencing effort were excluded.
methane, MT, and H2S concentrations were periodically measured in the headspace using a gas-tight syringe. The methane, MT, and H2S degradation rates were calculated from the slopes of concentration plotted versus time. The maximum methane oxidation rate (Vmax) and saturation constant (Km) were determined from the LineweavereBurk equation below (26): 1 Km 1 1 ¼ þ V Vmax ½S Vmax
(1)
where V represents the methane oxidation rate and [S] represents the methane concentration. The inhibition constants (Ki) of MT and H2S on methane oxidation were calculated using the equation for competitive inhibition, given by Williams et al. (27): 1 Km ¼ V Vmax
½i 1 1 þ 1þ ½S Vmax Ki
(2)
where [i] represents the MT or H2S concentration. All of the experiments were performed in triplicate. Inhibitory effects of methanethiol and hydrogen sulfide on methane oxidation by MMO pMMO was exclusively induced for the methane oxidation of the consortium by cultivating the culture in the presence of Cu. The bottles were prepared, as described above, with the addition of allylthiourea (SigmaeAldrich). Allylthiourea has been widely used as an inhibitor for pMMO (28,29). Allylthiourea was supplemented into the bottles at 50 mM of the final concentration. Methane, MT, and H2S gases were added to the bottles, for final concentrations of 68, 327, and 196.2 mmol L1 of liquid, respectively. The serum bottles were incubated at 30 C with 200 rpm of an agitation. The concentrations of methane, MT, and H2S in the headspace were monitored every 4 or 5 h using a gas-tight syringe. All of the experiments were performed in triplicate. Quantitative analysis of the pmoA gene expression One milliliter of each culture was collected from the serum bottles of the kinetic experiments earlier at 48, 12, and 5 experimental hours, which marked the end of the degradation periods for CH4, MT, and H2S, respectively (Figs. 2 and 3). The samples were immediately frozen at 70 C prior to analysis. Genomic DNA was extracted from the collected samples in duplicate using a NucleoSpin Soil Kit (MachereyeNagel GmbH & Co. KG) as described above. The DNA was eluted in 100 mL of the elution buffer and stored at 20 C. RNA was extracted from the collected samples in duplicate. For RNA extraction, 1 ml of the culture was added to 2-mL microcentrifuge tubes containing 0.1-mm glass beads and 1-mm zirconia/silica beads (0.5 g each). This procedure was described in detail by Kim et al. (21). RNA pellets were suspended in 50 mL of 0.1% DEPC-treated water. RNA was immediately purified using a Qiagen RNeasy Mini kit (Qiagen Inc., Valencia, CA, USA) with a RNase-Free DNase set (Qiagen Inc.), as specified by the manufacturer. RNA extracts were quantified using a
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FIG. 2. The effect of methanethiol (MT) concentrations on methane oxidation by the methane-oxidizing consortium. Time profiles of methane (a) and MT concentration (b). Closed and open symbols represent methane and MT, respectively. Closed circles, methane alone (68 mM); triangles, methane (68 mM) þ MT (163.5 mM); squares, methane (68 mM) þ MT (327 mM); inverted triangles, methane (68 mM) þ MT (490.5 mM); diamonds, methane (68 mM) þ MT (654 mM); crosses, control (no inoculation).
spectrophotometer. All RNAs were stored at 70 C prior to use. DNA contamination was confirmed by PCR, using the 340F-805R primer set. The PCR mixture and operating conditions have previously been outlined by Kim et al. (21). The PCR results were checked using a 2% agarose gel electrophoresis. No positive results were observed from the RNA extracts (data not shown). The RNA was reverse-transcribed using a Qiagen Omniscript RT kit (Qiagen), according to the manufacturer’s recommendations. The 20-mL reaction mixture contained 1 RT Sensiscript buffer, dNTP at a concentration of 0.5 mM, a random hexamer at 10 mM, 10 U of RNase inhibitor (RNaseOUT, Invitrogen Inc., Grand Island, USA), 1 mL of RT-Sensiscript reverse transcriptase, and 10 mL of template RNA. q-PCR was employed to quantify the pmoA gene expression of the methanotrophs using the primer set of A189f and mb661r, which targets the pmoA gene (30). The pmoA gene of Methylobacter luteus (NCIMB11914) was used to establish a standard curve for the quantitative detection. The PCR mixture and operating conditions were previously described (20). DNA and cDNA were used for the q-PCR templates in order to determine the pmoA copy numbers of the target DNA and mRNA transcript. The q-PCR was performed in five replicates. Gas analysis Methane was measured using gas chromatography (GC, 6850N, Agilent Technologies Inc., Santa Clara, CA, USA) equipped with a flame ionization detector (FID) and a wax column (30 m 0.32 mm 0.25 mm, Supelco Inc., Bellefonte, PA, USA), as previously described (20). The oven, injector, and detector temperatures were 100 C, 230 C, and 230 C, respectively. MT and H2S were analyzed using a GC (7890A, Agilent Technologies Inc., Santa Clara, USA) equipped with a flame photometric detector (FPD) and a DB-1 column (30 m 0.3 mm, J&W Scientific, Folsom, CA, USA). The oven, injector, and detector temperatures were 130 C, 200 C, and 250 C, respectively. The detection limits were 10, 3, and 3 ppm for CH4, MT, and H2S, respectively. Statistical analysis A statistical analysis of variance (ANOVA) was performed to validate the q-PCR abundance of each group using the SPSS 20 for Windows (IBM Corp., Armonk, NY, USA) program. A regression analysis was employed to estimate
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FIG. 3. The effect of hydrogen sulfide (H2S) concentrations on methane oxidation by the methane-oxidizing consortium. Time profiles of methane (a) and H2S concentration (b). Closed and open symbols represent methane and H2S, respectively. Closed circles, methane alone (68 mM); triangles, methane (68 mM) þ H2S (98.1 mM); squares, methane (68 mM) þ H2S (196.2 mM); inverted triangles, methane (68 mM) þ H2S (294.3 mM); diamonds, methane (68 mM) þ H2S (392.4 mM); crosses, control (no inoculation).
the relationship between the methane concentration and the methane oxidation rate. The level of significance was p < 0.05 for all of the statistical analyses.
RESULTS A bacterial community structure of a methane-oxidizing consortium The bacterial community of the methane-oxidizing consortium was analyzed using pyrosequencing assay. As shown in Fig. 1, 12 bacterial genera (Steroidobacter, Sphingopyxis, Opitutus, Methylocystis, unclassified Methylocystaceae, Meiothermus, Hyphomicrobium, unclassified Gammaproteobacteria, unclassified Chitinophagaceae, unclassified Bradyrhizobiaceae, Bosea, and unclassified Beijerinckiaceae) were detected in the consortium. The genus Methylocystis dominated the bacterial community in the consortium, followed by unclassified Chitinophagaceae, unclassified Gammaproteobacteria, Meiothermus, and Sphingopyxis. Methylocystis comprised 51 0.2% of the bacterial community in the methane-oxidizing consortium. Kinetic analysis for methanethiol and hydrogen sulfide effects on methane oxidation Fig. 2 shows the methane oxidation and MT degradation in the consortium. The consortium was capable of oxidizing methane without a lag period at a methane concentration of 68 mM in the absence of MT. However, methane oxidation was delayed in the presence of MT, and no methane oxidation was observed during an experimental period
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Methanethiol concentration in headspace (ppm)
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0
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FIG. 4. The effect of methanethiol (MT) and hydrogen sulfide (H2S) on the methane oxidation rates by a methane-oxidizing consortium at different methane concentrations.
at the MT concentrations above 490.5 mM (Fig. 2a). The methane oxidation rates decreased with increasing MT concentrations, from 1134.0 9.6 to 773.2 29.2 mmol g-dry cell weight (DCW)1 h1 at the MT concentrations of 0e327 mM. On the other hand, MT was degraded immediately by the methaneoxidizing consortium, and the degradation occurred even at a high concentration of 654 mM of MT without inhibition (Fig. 2b). The MT degradation rates showed 13.6e4.8 mmol g-DCW1 h1 at 327e654 mM of MT, which decreased as MT concentrations increased. The addition of H2S inhibited methane oxidation of the methane-oxidizing consortium (Fig. 3). The period for methane oxidation increased with increasing H2S concentrations, for instance, it took 4 days to oxidize approximately 90% of the methane in the presence of 392.4 mM H2S (Fig. 3a). The methane oxidation rates decreased from 1085.7 8.6 to 734.2 18.2 mmol gDCW1 h1 at the H2S concentrations of 0e196.2 mM. The consortium degraded H2S at concentrations of 98.1e392.4 mM without any delays. The H2S degradation rates ranged from 17.5 to 81.0 mmol g-DCW1 h1 at 98.1e392.4 mM of H2S, which were greater than the rates of MT degradation (Figs. 2b and 3b). H2S was completely degraded within 5 h, even at a high concentration of 392.4 mM (Fig. 3b). Fig. 4 shows the effect of MT and H2S on methane oxidation at different concentrations of methane. The methane oxidation rates of the consortium were measured at different methane concentrations, ranging from 27.2 to 204 mM. Then, they were compared with the methane oxidation rates which were measured in the presence of MT (327 mM) or H2S (196.2 mM). MT addition lead to a decrease of 13e31% for the methane oxidation, compared to the methane oxidation rates at 27.2e136 mM of methane alone. H2S addition caused a 31e38% decrease in the methane oxidation rates, compared to the rates at 27.2e136 mM of methane alone. However, no significant decrease of methane oxidation rates was observed at 204 mM of methane under the coexistence of MT or H2S. LineweavereBurk plots (Eq. 1) were constructed to estimate the Vmax, Km, and Ki values, based on MichaeliseMenten kinetics (data not shown). The methane-oxidizing consortium showed 3.7 mmol g-DCW1 h1 of Vmax and 184.1 mM of Km. The Ki values were determined from Eq. 2 for competitive inhibition (27), since it showed similar Vmax values, irrespective of MT and H2S addition. Ki parameters were calculated using Eq. 2 with the aforementioned
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Time (h) FIG. 5. Effects of allylthiourea on the degradation of methanethiol (MT) and hydrogen sulfide (H2S) by a methane-oxidizing consortium. Closed and open symbols represent MT or H2S degradation in the absence and presence of allylthiourea, respectively. (a) Time profile of MT concentration. Circles, MT alone (327 mM); squares, MT (327 mM) þ methane (68 mM); closed inverted triangle, control (no inoculation). (b) Time profile of H2S concentration. Circles, H2S alone (196.2 mM); squares, H2S (196.2 mM) þ methane (68 mM); closed inverted triangle, control (no inoculation).
values of Vmax and Km. The Ki values of MT and H2S were 1504.8 and 359.8 mM, respectively. Inhibitory effects of methanethiol and hydrogen sulfide on methane oxidation by MMO The MMO assay was performed to investigate the effects of MT and H2S on methane oxidation by the methane-oxidizing consortium. Particularly, pMMO was targeted for the assay, since the consortium was induced to express pMMO exclusively by adding Cu ions into the media during cultivation. Allylthiourea is a well-known inhibitor on the synthesis of pMMO (28,29). Yu et al. (29) reported that allylthiourea addition reduced the amount of intracellular copper by preventing entry of copper into the cells. This presumably leads to the decrease of the methanotrophic pMMO expression. No methane oxidation occurred in the presence of allylthiourea (data not shown). This indicated that methane was oxidized exclusively by the pMMO of the methane-oxidizing consortium. The MT was completely degraded within 12 h in the absence of
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FIG. 6. Effects of methanethiol (MT) and hydrogen sulfide (H2S) on the pmoA gene expression in the methane-oxidizing consortium. The pmoA gene levels were analyzed using cultures that were sampled from the serum bottles of the kinetic experiments (Figs. 2 and 3). In the coexistence of methane with MT and H2S, the culture was collected after 5, 12, 48 h of incubation time (shown in Figs. 2 and 3), which are ends of degradation periods for H2S, MT, and CH4, respectively. The initial concentrations of CH4, MT, and H2S were 68, 327, and 196.2 mM respectively. (a) Abundance of pmoA gene at DNA level. (b) Abundance of pmoA gene at mRNA transcript level.
allylthiourea (Fig. 5a). However, MT degradation was retarded in the presence of allylthiourea and only 41e44% of the MT was removed for 12 h. H2S was degraded by the consortium, irrespective of allylthiourea addition, showing a slight increase in the presence of allylthiourea (Fig. 5b). H2S was degraded 99.6 0.5% in the presence of allylthiourea within 6.5 h, while it was 89.0 1.9% in the absence of allylthiourea. Quantification of pmoA gene expression in the methaneoxidizing consortium q-PCR was performed to quantify the
pmoA gene copy numbers in the methane-oxidizing consortium using DNA and RNA extracts from the kinetic experiments. The initial DNA and mRNA transcript levels of the pmoA gene were 9.9 1.2 106 and 1.6 0.7 105 pmoA gene copy number gDCW1, respectively (Fig. 6). The DNA levels of the pmoA gene decreased to 2.6 0.2 106 and 3.5 0.2 106 pmoA gene copy number g-DCW1 after its degradation for MT or H2S alone, respectively, whereas it was slightly increased to 1.6 0.1 107 pmoA gene copy number g-DCW1 after methane oxidation for methane alone (Fig. 6a). In the coexistence of methane with MT and H2S, the DNA levels of the pmoA gene decreased to 5.4 0.32 106 and 2.1 1.0 106 pmoA gene copy number gDCW1 after MT and H2S degradation (at the point of 12 and 5 h in incubation time), respectively, methane oxidation recovered to the initial DNA level of w107 pmoA gene copy number g-DCW1 (at the point of 48 h in incubation time). No statistical differences were observed among the DNA levels of the pmoA gene as compared to the initial value. In contrast, the amount of mRNA transcript for the pmoA gene expression changed significantly between the samples (Fig. 6b). For example, in the coexistence of methane with MT and H2S, the mRNA transcript levels of the pmoA gene decreased greatly to 3.8 4.2 103 and 6.6 1.2 103 pmoA gene copy number g-DCW1 after MT and H2S degradation (at the point of 12 and 5 h in incubation time), respectively, compared to the initial level, and they were subsequently recovered to 2.2e6.6 0.7 105 pmoA gene copy number g-DCW1 after methane oxidation (at the point of 48 h in incubation time). The decrease and recovery of the mRNA transcript levels of the pmoA gene were statistically proven using ANOVA analysis.
DISCUSSION Kinetic and enzymatic approaches were performed to investigate the effects of MT and H2S on methane oxidation by the methane-oxidizing consortium. The consortium obtained in this study oxidized methane successfully without a lag period, since the methanotrophs were actively cultured by continuously enriching the consortium with methane as the sole carbon and energy source (Figs. 2 and 3). Methanotrophs are classified into two major groups of type I and type II, depending on the DNA content, intracellular membrane arrangement, carbon assimilation pathway, and PLFA composition (31). In this study, the type II methanotroph Methylocystis dominated the bacterial community, comprising 51 0.2% of the bacterial community in the methane-oxidizing consortium (Fig. 1). Generally, the methane concentration can affect methanotrophic community compositions (32,33). The consortium was continuously exposed to a relatively high methane concentration of 68 mM during enrichment. This condition supplied a favorable environment to type II methanotrophs and this could lead to the dominance of Methylocystis in the bacterial community. Our observations support previous studies, which reported that type II methanotrophs preferred methane rich environments and outcompeted type I methanotrophs in high methane concentrations (32,34). Methylocystis is a well-known methane-oxidizing bacterium (8), indicating that this genus plays a primary role in methane oxidation. The MMO assay revealed that the methane was oxidized exclusively by pMMO (data not shown). This is not surprising, as the consortium was continuously cultured in an NMS medium supplemented with Cu ions. Generally, Cu regulates MMO expression in methanotrophs and the pMMO is expressed at high concentrations of Cu >0.8 mmol of Cu per g-DCW (8). The consortium showed 3.7 mmol g-DCW1 h1 of Vmax and 184.1 mM of Km. The Vmax and Km values are greater than or comparable to the levels observed in previous studies. For instance,
VOL. 120, 2015 Choi et al. (35) reported the Vmax and Km values for methane oxidation by a mixed culture of methanotrophs were 326.8 mmol of CH4 g-DCW1 h1 and 143.8 mM, respectively. Lee et al. (17) calculated Vmax and Km values for methane oxidation by Methylocystis sp. M6 as 4.93 mmol of CH4 g-DCW1 h1 and 23 mM, respectively. Kinetic analysis revealed that MT and H2S had inhibitory effects on methane oxidation by the consortium (Figs. 2 and 3). Methane oxidation was delayed in the presence of MT or H2S, and their methane oxidation rates decreased to 424.4e773.2 and 330.0e484.3 mmol g-DCW1 h1, respectively, corresponding to a 13e31 and 30e50% decrease of the oxidation rates at 27.2e68 mM of methane alone (Figs. 2e4). This is consistent with earlier studies, which reported that the methane oxidation of Methylocysitis sp. M6 was inhibited by the presence of MT and H2S (17). Börjesson (36) studied the inhibition of methane oxidation by volatile sulfur compounds in landfill cover soils, describing MT’s inhibitory effects on methane oxidation rates. Jäckel et al. (37) found that the addition of MT resulted in an inhibition of methane oxidation in deciduous forest soil. Cáceres et al. (16) reported that H2S over 0.05% (v/v) decreased the methane oxidation rate of Methylocystis sp. The expression of pMMO in the consortium was quantified using a q-PCR with DNA and RNA extracts from the kinetic experiments. The results of the pmoA gene expression also support the kinetic analysis (Fig. 6). In the coexistence of methane with MT and H2S, the DNA and mRNA transcript levels for the pmoA gene expression were decreased to 2.1e5.4 106 and 3.8e6.6 103 pmoA gene copy number g-DCW1 after MT (at the point of 12 h in incubation time) and H2S degradation (at the point of 5 h in incubation time), respectively, as compared to their initial levels. However, both the amount of DNA and mRNA transcript for the pmoA gene were recovered to the initial levels of w107 and 105 pmoA gene copy number g-DCW1 after methane oxidation (at the point of 48 h in incubation time), respectively. Interestingly, the changes in the pmoA gene expression differed between the results of the DNA- and RNA-based analyses. Although the DNA levels of the pmoA gene were decreased after MT and H2S degradation, the differences were observed on the same order of 106 level, whereas the mRNA transcripts showed a 100-fold decrease. These results were confirmed by a statistical analysis. No difference was observed between the DNA levels of the pmoA gene; however, the mRNA transcripts levels varied significantly. The analysis of DNA-based gene expression can be used for evaluation of microbial activities and microbial ecology, however that provides relatively little information since DNA is known to be stable even in dead cells (38). On the other hand, the analysis of mRNA-based gene expression can provide relatively more information on the microbial activities and ecological significance as it has short half-life (wminutes) which is primary regulated at the level of transcription (39e41). For instance, Dumont et al. (42) found that the C13 labelling of the mRNA for the pmoA gene was quicker than DNA, following the incubation of lake sediment with 13CH4, demonstrating that mRNA-stable isotope probing (SIP) is more sensitive than DNA-SIP. Han and Semrau (43) examined the quantification of gene expression in methanotrophs using a competitive reverse transcription-polymerase chain reaction and reported that the amount of mRNA transcript for the pmoA gene correlated well with the whole-cell pMMO activity. In this study, the expression of pmoA gene was quantified at the point of 5, 12, and 48 h in incubation time of the kinetic experiments, this presumably only induced the changes of the mRNA-based pmoA gene levels because of short time of incubation. Based on the previous reports and our results, it is possible to conclude that MT and H2S dramatically inhibit the pMMO activity of the consortium since mRNA levels were decreased due to MT and H2S addition, resulting in a decrease of methane oxidation by repressing pmoA gene expression. A comparison between the results of DNA- and RNA-
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based pmoA gene levels demonstrated that the DNA results alone can lead to an underestimation of the active gene expression since it is probably incapable of sensitively reflecting changes of pMMO activity. The LineweavereBurk plots revealed that MT and H2S follow a typical competitive inhibition model, since they had the same yintercepts as those of methane alone (data not shown). As shown in Fig. 4, the methane oxidation rates were similar at 204 mM of methane, regardless of whether MT or H2S were added. This means that if the amount of methane is very high, the effect of the inhibitors (MT and H2S) becomes less noticeable. The inhibition constants of MT and H2S were 1504.8 and 359.8 mM, respectively, determined using Eq. 2 for competitive inhibition. These values indicate that H2S has greater inhibitory effects on methane oxidation than MT. This is consistent with an earlier study for the inhibitory effects of MT on methane oxidation, which followed a competitive inhibition model with an inhibition constant of 40.6 18.5 nmol or 1.95 mg MT g-dry soil1 (36). Although MT and H2S inhibited methane oxidation by the methane-oxidizing consortium, methane oxidation occurred in the presence of MT (327 mM) and H2S (196.2 mM). This suggests that MT and H2S are possibly classified as reversible inhibitors. The results of the mRNA transcript levels for the pmoA gene expression also support this observation, which were recovered to the level of 105 pmoA gene copy number g-DCW1 after methane oxidation (Fig. 6). The methane-oxidizing consortium was capable of degrading MT and H2S, and MT and H2S degradation occurred without a lag period (Figs. 2, 3, 5, and 6). MT degradation rapidly decreased in the presence of allylthiourea, indicating that most of the MT was degraded by pMMO (Fig. 5a). This suggests that methanotrophs play an important role in MT degradation. In addition, MT was slightly degraded in the presence of allylthiourea, compared to the abiotic experiment (control), suggesting that other microorganisms or enzymes were marginally involved in the MT degradation as well. Conversely, H2S degradation occurred, irrespective of allylthiourea addition, indicating that pMMO was not associated with the H2S degradation (Fig. 5b). Interestingly, H2S was completely degraded by the consortium, suggesting that other microorganisms or enzymes play a primary role in the H2S degradation. Thus, we can conclude that methanotrophs are capable of cometabolizing MT, but not H2S, while other microorganisms or enzymes are primary involved in the H2S degradation. Börjesson (36) observed that MT was rapidly consumed, with a rate of 16.03 3.6 mmol MT g-dry soil1 h1, in landfill cover soils. Zhang et al. (44) reported that Hyphomicrobium sp. is capable of removing MT and H2S. Lee et al. (45) found that Sphingopyxis sp. degrades MT and H2S. These studies indicate that the genera Hyphomicrobium and Sphingopyxis have potential for MT and H2S degradation in the consortium (Fig. 1). The objective of this study was to investigate the kinetic and enzymatic inhibitory effects of MT and H2S on methane oxidation of the consortium. MT and H2S showed competitive inhibition on methane oxidation. MT was primary degraded by the cometabolism of pMMO, whereas H2S was degraded by other microorganisms or enzymes in the consortium. Our findings demonstrated that pMMO is considerable for the cometabolism of MT, despite of its narrow range of substrate specificity than sMMO, since the pMMO was easily reproducible (as shown in the analysis of pmoA gene expression). On the basis of the results, we conclude that this study provides valuable information for methane mitigation and its kinetics and interactions with sulfur compounds. ACKNOWLEDGMENTS This research was supported by the Basic Science Research Program through the National Research Foundation of Korea (NRF),
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funded by the Ministry of Science, ICT and Future Planning (NRF2012R1A2A2A03046724), and by the National Research Foundation of Korea (NRF 2009-83527). E. H. Lee was financially supported by National Research Foundation of Korea (NRF-2014R1A1A3051952). References 1. Solomon, S.: Climate change 2007-the physical science basis: Working group I contribution to the fourth assessment report of the IPCC. Cambridge University Press (2007). 2. Smet, E. and Van Langenhove, H.: Abatement of volatile organic sulfur compounds in odorous emissions from the bio-industry, Biodegradation, 9, 273e284 (1998). 3. Themelis, N. J. and Ulloa, P. A.: Methane generation in landfills, Renew. Energy, 32, 1243e1257 (2007). 4. Saral, A., Demir, S., and Yildiz, S.: Assessment of odorous VOCs released from a main MSW landfill site in Istanbul-Turkey via a modelling approach, J. Hazard. Mater., 168, 338e345 (2009). 5. Gendebien, A., Pauwels, M., Constant, M., Ledrut-Damanet, M.-J., Nyns, E.-J., Willumsen, H.-C., Butson, J., Fabry, R., and Ferrero, G.-L.: Landfill gas. From environment to energy. Commission of the European Communities, Luxembourg (1992). 6. U.S. Environmental Protection Agency: Municipal solid waste landfills, section 2.4, in: AP-42, 5th ed. Compilation of air pollutant emissions factors. Volume I: Stationary point and area sources. U.S. Environmental Protection Agency, Research Triangle Park, NC (1995). 7. Lassey, K. R., Etheridge, D. M., Lowe, D. C., Smith, A. M., and Ferretti, D. F.: Centennial evolution of the atmospheric methane budget: what do the carbon isotopes tell us? Atmos. Chem. Phys., 7, 2119e2139 (2007). 8. Hanson, R. S. and Hanson, T. E.: Methanotrophic bacteria, Microbiol. Rev., 60, 439e471 (1996). 9. Vorobev, A. V., Baani, M., Doronina, N. V., Brady, A. L., Liesack, W., Dunfield, P. F., and Dedysh, S. N.: Methyloferula stellata gen. nov., sp. nov., an acidophilic, obligately methanotrophic bacterium that possesses only a soluble methane monooxygenase, Int. J. Syst. Evol. Microbiol., 61, 2456e2463 (2011). 10. McDonald, I. R., Bodrossy, L., Chen, Y., and Murrell, J. C.: Molecular ecology techniques for the study of aerobic methanotrophs, Appl. Environ. Microbiol., 74, 1305e1315 (2008). 11. Murrell, J. C., McDonald, I. R., and Gilbert, B.: Regulation of expression of methane monooxygenases by copper ions, Trends Microbiol., 8, 221e225 (2000). 12. Theisen, A. R., Ali, M. H., Radajewski, S., Dumont, M. G., Dunfield, P. F., McDonald, I. R., Dedysh, S. N., Miguez, C. B., and Murrell, J. C.: Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2, Mol. Microbiol., 58, 682e692 (2005). 13. Hakemian, A. S. and Rosenzweig, A. C.: The biochemistry of methane oxidation,, Annu. Rev. Biochem., 76, 223e241 (2007). 14. Semrau, J. D., DiSpirito, A. A., and Yoon, S.: Methanotrophs and copper, FEMS Microbiol. Rev., 34, 496e531 (2010). 15. Morton, J. D., Hayes, K. F., and Semrau, J. D.: Effect of copper speciation on whole-cell soluble methane monooxygenase activity in Methylosinus trichosporium OB3b, Appl. Environ. Microbiol., 66, 1730e1733 (2000). 16. Caceres, M., Gentina, J. C., and Aroca, G.: Oxidation of methane by Methylomicrobium album and Methylocystis sp. in the presence of H2S and NH3, Biotechnol. Lett., 36, 69e74 (2014). 17. Lee, E. H., Yi, T., Moon, K. E., Park, H., Ryu, H. W., and Cho, K. S.: Characterization of methane oxidation by a methanotroph isolated from a landfill cover soil, South Korea, J. Microbiol. Biotechnol., 21, 753e756 (2011). 18. Colby, J., Stirling, D. I., and Dalton, H.: The soluble methane mono-oxygenase of Methylococcus capsulatus (Bath). Its ability to oxygenate n-alkanes, n-alkenes, ethers, and alicyclic, aromatic and heterocyclic compounds, Biochem. J., 165, 395e402 (1977). 19. Whittenbury, R. and Dalton, H.: The methylotrophic bacteria, pp. 894e902, in: Starr, M., Stolp, H., Trüper, H., Balows, A., and Schlegel, H. (Eds.), The prokaryotes. Springer Berlin Heidelberg (1981). 20. Lee, E. H., Park, H., and Cho, K. S.: Characterization of methane, benzene and toluene-oxidizing consortia enriched from landfill and riparian wetland soils, J. Hazard. Mater., 184, 313e320 (2010). 21. Kim, T. G., Moon, K. E., Yun, J., and Cho, K. S.: Comparison of RNA- and DNAbased bacterial communities in a lab-scale methane-degrading biocover, Appl. Microbiol. Biotechnol., 97, 3171e3181 (2013).
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