Shifts in Identity and Activity of Methanotrophs in Arctic Lake Sediments in Response to Temperature Changes
ABSTRACT
Methane (CH4) flux to the atmosphere is mitigated via microbial CH4 oxidation in sediments and water. As arctic temperatures increase, understanding the effects of temperature on the activity and identity of methanotrophs in arctic lake sediments is important to predicting future CH4 emissions. We used DNA-based stable-isotope probing (SIP), quantitative PCR (Q-PCR), and pyrosequencing analyses to identify and characterize methanotrophic communities active at a range of temperatures (4°C, 10°C, and 21°C) in sediments (to a depth of 25 cm) sampled from Lake Qalluuraq on the North Slope of Alaska. CH4 oxidation activity was measured in microcosm incubations containing sediments at all temperatures, with the highest CH4 oxidation potential of 37.5 μmol g−1 day−1 in the uppermost (depth, 0 to 1 cm) sediment at 21°C after 2 to 5 days of incubation. Q-PCR of pmoA and of the 16S rRNA genes of type I and type II methanotrophs, and pyrosequencing of 16S rRNA genes in 13C-labeled DNA obtained by SIP demonstrated that the type I methanotrophs Methylobacter, Methylomonas, and Methylosoma dominated carbon acquisition from CH4 in the sediments. The identity and relative abundance of active methanotrophs differed with the incubation temperature. Methylotrophs were also abundant in the microbial community that derived carbon from CH4, especially in the deeper sediments (depth, 15 to 20 cm) at low temperatures (4°C and 10°C), and showed a good linear relationship (R = 0.82) with the relative abundances of methanotrophs in pyrosequencing reads. This study describes for the first time how methanotrophic communities in arctic lake sediments respond to temperature variations.
INTRODUCTION
Climate change is an important issue, with some of the most marked changes occurring in the Arctic (63). The mean annual temperature of permafrost in some areas of the Arctic has recently increased (59), and continued warming and thawing are predicted (36). Freshwater thermokarst lakes can form in topographic depressions as permafrost melts and are widely distributed in the Arctic, contributing significantly to the atmospheric methane (CH4) budget (70, 71). CH4 is an important greenhouse gas that is about 23 times more potent than carbon dioxide over a 100-year period (31). Increased thawing of permafrost as a result of warming in the Arctic could act as a positive feedback to climate warming through the release of carbon previously sequestered in permafrost (1, 60) as CH4 emissions from arctic lakes (1, 70). If the frozen Pleistocene-age carbon in the Arctic were to be released during the next 500 to 1,000 years, the average CH4 emission rate from lakes would be approximately 50 to 100 Tg year−1 (70), which is 8 to 17% of the contemporary atmospheric CH4 budget.
The net flux of CH4 to the atmosphere is mitigated to some extent by CH4 oxidation within the water column and sediments (41, 42, 67). Microbial CH4 oxidation occurs aerobically and anaerobically. At the sediment-water interface in freshwater ecosystems, where CH4 and oxygen gradients overlap, aerobic CH4 oxidation is an important process in reducing CH4 release to the atmosphere (4, 10, 21, 27).
Methanotrophic community structure and activity differ with temperature (12, 41, 49) as well as with other environmental conditions, such as the concentrations of CH4 and O2, pH, and nitrogen stress (27, 61). Temperature also influences the enzymatic activity of methanotrophs, including hydroxypyruvate reductase, hexulose phosphate synthase, formate dehydrogenase, and ribulose bisphosphate carboxylase activities (45). Most methanotrophs available in pure culture are mesophiles, although psychrophilic methylotrophs have been described (7, 51). Although active type I and type II methanotrophs have been found over a wide range of temperatures (67), in general, type II methanotrophs are favored at temperatures of >15°C (47, 68) while type I methanotrophs prevail at low temperatures (0 to 10°C) (6, 26, 41, 69, 72, 73). However, type II methanotrophs—Methylocella species—have been found to predominate in acidic arctic tundra soils (5, 20). Some mesophilic methanotrophs can maintain viability with exposure to subzero temperatures, while psychrophilic and psychrotolerant methanotrophs can maintain growth and/or activity at low temperatures (67).
In the Arctic, ice on shallow ponds and lakes generally begins to melt in June and is absent between June and September (46). The average temperatures at the surfaces of the sediments in some shallow ponds and lakes in the Arctic have been shown to be very close to the average water temperatures. The mean daily water temperature is usually below 4°C but rises rapidly in June, with a seasonal peak at about 16 to 20°C in the first week of July (9, 18, 46). Moreover, most shallow arctic lakes are thermokarst lakes (70) and would be expected to undergo complete drainage once the underlying permafrost has thawed, enabling permeation of water downward through the thaw bulbs (34). At that time, methanotrophs in the shallow, drained sediments in a drained landscape would have direct exposure to climate perturbations and greater depth of O2 penetration. Because of the crucial role of methanotrophic activity in controlling CH4 emissions (27, 61), describing how the activity and structure of methanotrophic communities respond to temperature fluctuations in Arctic lake sediments is important for predicting how future climate warming and permafrost thaw will influence CH4 emissions from Arctic lakes. However, the response of active methanotrophs to temperature changes in arctic lakes has not been well studied, although these lakes are significant contributors to atmospheric CH4 (70, 71).
We employed DNA-based stable-isotope probing (SIP), quantitative PCR (Q-PCR), and pyrosequencing analyses to identify and characterize the metabolically active methanotrophs in arctic lake sediments along a 25-cm sediment core sectioned at intervals of 0 to 1, 1 to 3, 3 to 5, 5 to 10, 10 to 15, 15 to 20, and 20 to 25 cm. We also tracked the activity and community structure of the active methanotrophs in sediments in response to three incubation temperatures (4°C, 10°C, and 21°C), representing a range above and below the peak annual temperatures recorded for some arctic lake sediments (about 16 to 20°C) (46).
MATERIALS AND METHODS
Study site and sampling.
The study site, Lake Qalluuraq, is located on the North Slope of the Brooks Range in Alaska, about 136 m away from a site with continuous CH4 ebullition (70°22.669′N, 157°20.925′W) (75). The depth, temperature, and dissolved oxygen (DO) concentration of water were about 170 cm, 16 to 17°C, and 10 mg liter−1, respectively, when sampling was conducted in July 2009. Core tubes (length, 2.5 m; diameter, 7 cm) were used for the collection of sediment samples in depth increments of 1 to 2 cm within the top 5 cm of sediment and then every 5 cm for the remaining length of the sediment core. The sediment subsamples were immediately placed in plastic zipper freezer bags and were homogenized; then subsamples were frozen at −80°C for later molecular studies. Sediment samples in the plastic bags were kept at 4°C for CH4 oxidation incubations and SIP experiments (described below). The particle composition of the experimental sediment mixture (depth, 0 to 25 cm) was as follows: at >2 mm, 0.6%; at 0.05 to 0.1 mm, 67.2%; at 0.05 to 0.002 mm, 22.2%; at <0.002 mm, 10%. The total carbon and total nitrogen contents of the sediments, measured every 5 cm, were 14.8% ± 3.1% and 0.9% ± 0.2%, respectively (means ± standard deviations for 5 samples).
For measurement of dissolved CH4 concentrations, sediments were collected as 3-ml plugs at 2-cm intervals from a 20-cm core. The samples were transferred to 20-ml serum vials, sealed with 1-cm-thick septa, and stored at −20°C. The headspace hydrocarbon concentrations were in the vials was analyzed by gas chromatography-flame ionization detection (GC-FID) (52). Headspace concentrations were converted to aqueous concentrations by using the method of Hoehler et al. (29).
SIP microcosms and measurement of CH4 oxidation potential.
Incubations for SIP and for measurement of CH4 oxidation potential were conducted in microcosms. Sediment microcosms were constructed by adding approximately 5 g (wet weight) of each subsample of the sediments to a sterile 60-ml glass serum vial that was crimp-sealed with a butyl rubber stopper. Each microcosm was injected with 13CH4 (99 atom% 13C; Sigma-Aldrich) or CH4 (99.5% pure, as a control) to 10% (vol/vol). Calculated by Henry's law and the van't Hoff equation to correct for the effect of temperature on CH4 solubility (39), the theoretical saturated CH4 concentration in the pore water of microcosm sediments was 151 μmol liter−1 at the initial CH4 concentration of 10% (vol/vol). In the shallow ponds and lakes in the Arctic, the mean daily water temperature is usually below 4°C but rises rapidly in June and experiences a seasonal peak at about 16 to 20°C in the first week of July (9, 18, 46). Considering the seasonal temperatures of the arctic lakes, microcosms were incubated at 4°C, 10°C, or 21°C on a shaker (100 rpm). All treatments were performed in triplicate. The disappearance of CH4 from the headspace was monitored using gas chromatography. Gas headspace samples (50 μl) were withdrawn from microcosms at intervals and were analyzed for residual CH4 by using GC-FID during incubation.
CH4 oxidation potential was measured at incubation times of 2 days, 5 days, and 10 days at 21°C, 4 days, 10 days, and 20 days at 10°C, and 5 days, 15 days, and 30 days at 4°C; assessed from the zero-order decrease in CH4 concentration in the headspace of the serum vials within 12 to 72 h (35); and expressed as micromoles of CH4 per gram (dry weight) per day. For SIP, after >90% of the initial CH4 concentration had been oxidized, the serum vials were flushed with air, and the initial CH4 concentration (10% [vol/vol]) was reestablished. The incubation was continued in the same manner until approximately 0.2 mmol 13CH4 or CH4 g (wet weight)−1 was consumed (55 to 74 days at 21°C, 144 to 212 days at 10°C, and 212 to 248 days at 4°C), quantities reportedly required to obtain a visible 13C-labeled DNA band following ultracentrifugation of DNA from 1 g of soil (56). Sediment samples were then harvested and were frozen immediately at −80°C.
DNA extraction and separation of 13C-labeled DNA.
DNA was extracted from about 0.5 g of the sediment subsamples subjected to SIP by using the Bio 101 Fast DNA Spin kit for soil (MP Biomedicals, Solon, OH). The equilibrium (isopycnic) density gradient centrifugation and fractionation procedures were modified from the method described by Leigh et al. (38) and used a cesium trifluoroacetate (CsTFA; GE Healthcare, United Kingdom) solution with a starting buoyant density (BD) of 1.60 g ml−1. Samples of 3 μg DNA were combined with the CsTFA solution and were loaded into Beckman polyallomer Quick-Seal centrifuge tubes (13 by 51 mm), which were then sealed and spun in an NVT 100 rotor in an Optima L-100 XP ultracentrifuge (Beckman Coulter) at 134,260 × g and 25°C for about 70 h. Gradients were fractionated into 250-μl fractions by displacement with nuclease-free water by using a syringe pump at a flow rate of 500 μl min−1. The BDs of fractions were measured gravimetrically by weighing aliquots of each fraction from a blank gradient, containing water in place of DNA, run in parallel. Sample DNA was precipitated from fractions with isopropanol overnight at −20°C; then pellets were washed twice with isopropanol and were resuspended in 20 μl nuclease-free water (Sigma-Aldrich).
The relative abundances of bacterial DNA in gradient fractions ranging in BD from 1.558 to 1.663 g ml−1 were determined by Q-PCR as described by Leigh et al. (38). After the range of fractions containing 13C-labeled DNA or unlabeled DNA was identified, they were combined to compose compiled “heavy” fractions from each sample for subsequent molecular analyses.
Q-PCR analysis.
Q-PCR of 16S rRNA genes of type I and type II methanotrophs and of the pmoA gene in the heavy fractions from 13C-labeled DNA and unlabeled control DNA was performed using primer sets U785F (3)/MethT1bR (74) and U785F/MethT2R (74) for 16S rRNA genes of type I and type II methanotrophs, respectively, and primer set A189F (30)/mb661R (17) for the pmoA gene. Q-PCR was conducted in 4 replicates of 15-μl reaction mixtures containing SYBR green master mix (Applied Biosystems, Foster City, CA), 4.5 pmol each primer, and 1 μl template. Thermal cycler conditions were as follows: an initial stage at 50°C for 2 min; denaturation at 95°C for 5 min; and 60 cycles of 95°C for 30 s, 58°C for 30 s, and either 72°C for 45 s for the pmoA gene or 72°C for 30 s for the 16S rRNA genes of type I and type II methanotrophs. Standards were made from 10-fold dilutions of linearized plasmids containing the same fragments of the 16S rRNA genes of type I and type II methanotrophs and the pmoA gene as the Q-PCR product that was cloned from amplified pure-culture DNA. The limit of detection by Q-PCR was about 102 copies per reaction for the pmoA gene and about 103 copies per reaction for the 16S rRNA genes of type I and type II methanotrophs.
16S rRNA gene sequence analysis.
Sequence analyses were performed on bar-coded 16S rRNA gene amplicons from 13C-labeled DNA by pyrosequencing. The primer set used consisted of 577F (5′-CGTATCGCCTCCCTCGCGCCATCAG [bar code] AYTGGGYDTAAAGNG-3′) and 926R (5′-CTATGCGCCTTGCCAGCCCGCTCAGCCGTCAATTCMTTTRAGT-3′) (with the sequencing adaptors in boldface). PCR was run using the same conditions as for terminal restriction fragment length polymorphism (T-RFLP) but with different primers. For each sample, 12 replicate 25-μl PCR mixtures were created to obtain abundant PCR products and to span variations in template and amplification bias. PCR products were then purified and concentrated using the QIAquick PCR purification kit (Qiagen Inc., Valencia, CA). Products were gel purified using the QIAquick gel extraction kit (Qiagen Inc., Valencia, CA), cleaned again with the QIAquick PCR purification kit, and eluted with 20 μl of EB buffer (10 mM Tris-Cl, pH 8.5). Clean products were quantified using the NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE) and were mixed in equal amounts (20 ng each). Pyrosequencing was performed using Roche 454 GS FLX Titanium sequencing (454 Life Sciences, Branford, CT) at the Research Technology Support Facility at Michigan State University (East Lansing, MI).
Sequences were first trimmed of the primer region, and low-quality sequences were removed by the pyrosequencing pipeline of the Ribosomal Database Project (RDP). In our libraries, 41% to 66% of the sequences in the libraries (sequence libraries ranged from 2,961 to 4,474 reads in size) passed this filter (average length, 330 to 331 bp) and were assigned to taxonomic groups by RDP's Naïve Bayesian Classifier (confidence threshold, 80%). The only exception was the library obtained from 13C-labeled DNA (sediment depth, 0 to 1 cm) at 10°C, in which 23% of the 3,102 total library sequences passed the filter.
Nucleotide sequence accession number.
The nucleotide sequences determined in this study have been deposited in the NCBI Short Read Archive under accession number SRP005485.
RESULTS
In situ CH4 concentrations.
Dissolved CH4 concentrations increase gradually from 1.5 μmol liter−1 at the sediment-water interface to 861.3 μmol liter−1 at a 19-cm sediment depth (Fig. 1). At the continuous CH4 ebullition site in the lake reported here, dissolved CH4 concentrations were less than 15 μmol liter−1 from a sediment depth of 0 to 10 cm and increased to 1,334 μmol liter−1 at a 25-cm sediment depth. In other parts of this lake, as well as in other lakes in Alaska in our study, dissolved CH4 concentrations at a sediment depth of 1.5 cm were 109 to 505 μmol liter−1, and the maximum measured dissolved CH4 concentration was 2,062 μmol liter−1 at a 25.5-cm sediment depth (28). The saturated CH4 concentration, calculated by Henry's law and the van't Hoff equation, in the pore water in SIP microcosms was 151 μmol liter−1 at the initial CH4 concentration of 10% (vol/vol). In comparison, the CH4 concentration in the pore water in the microcosms is within the range of ambient conditions.
Fig 1
CH4 oxidation potential.
CH4 oxidation potential measurement compares the relative activities of the methanotrophic populations among samples from different environmental conditions (35). CH4 oxidation potential was measured during SIP incubations conducted at 4°C, 10°C, and 21°C, with a headspace of CH4 at an initial concentration of 10% (vol/vol) in air. Measurable CH4 consumption was first detected after 2 days of incubation in the microcosms from a sediment depth of 0 to 10 cm incubated at 21°C. The highest CH4 oxidation potential, 37.5 μmol g−1 day−1, was recorded in sediments collected from a depth of 0 to 1 cm (Fig. 2). Although CH4 consumption was initially slow or undetectable in some microcosms (for example, in those from depths of 10 to 15 cm, 15 to 20 cm, and 20 to 25 cm), CH4 consumption was observed in all the sediment microcosms at 21°C after incubation for 5 days. After incubation at 21°C and 10°C for 10 days and 20 days, respectively, the sediments from the depth of 10 to 25 cm showed higher CH4 oxidation potentials, 47.2 to 53.6 μmol g−1 day−1 and 27.3 to 29.5 μmol g−1 day−1, than the upper sediments. The CH4 oxidation potentials observed increased as the incubation temperature increased from 4°C to 21°C. In the 0- to 5-cm sediment depth range, the CH4 oxidation potentials at 21°C (on day 5) were 0.6 to 1.6 times and 37.0 to 160.5 times greater than those at 10°C (on day 4) and 4°C (on day 5), respectively.
Fig 2
Identification of the CH4-utilizing bacterial community.
Q-PCR analyses of gradient fractions, indicating the relative abundance of bacterial DNA throughout the gradient, demonstrated that significant quantities of 13C-labeled DNA were present in the sediments incubated with 13CH4 (see Fig. S1 in the supplemental material). Unlabeled control DNA from the sediment at 21°C had a peak indicating abundant DNA at a BD of 1.600 g ml−1 (see Fig. S1d), and the DNA distributions in unlabeled controls at 4°C and 10°C were similar to those at 21°C (data not shown). Little or no bacterial DNA was detected in fractions with BDs higher than 1.619 g ml−1 for unlabeled control DNA, while a second peak was present in labeled DNA, indicating the presence of 13C-labeled DNA (see Fig. S1a, b, and c). Based on the Q-PCR results, we combined the fractions with BDs of 1.619 to 1.643 g ml−1 as compiled heavy fractions for each sample prior to downstream molecular analyses.
To characterize bacterial communities active in assimilating carbon from CH4, we selected 13C-labeled DNA samples from the uppermost sediment (depth, 0 to 1 cm) and from deeper sediments (15 to 20 cm) incubated at the three temperatures (i.e., 4°C, 10°C, and 21°C) for pyrosequencing analyses. Because the pmoA genes (encoding a subunit of particulate methane monooxygenase) of some novel aerobic methanotrophs, including Crenothrix polyspora, and of members of the phylum Verrucomicrobia are not detectable by use of standard primers in PCR amplification (22, 33, 53, 66) and no pmoA gene is present in Methylocella species (19), we relied on bacterial 16S rRNA gene sequencing instead of pmoA genes for 13C-labeled DNA pyrosequencing analysis to ensure that we captured the diversity of active methanotrophs. Although the numbers of pyrosequencing reads ranged from 699 to 2,440 in the uppermost sediment and from 1,782 to 2,934 in the deep sediment (Table 1), rarefaction curves showed a higher diversity in the uppermost sediment than in the deep sediment (Fig. 3). The relative abundances of different phylogenetic groups present in 13C-labeled DNA were determined based on the frequency of pyrosequencing reads associated with different taxa as determined using the RDP classifier (Table 1). The phylum most frequently detected in 13C-labeled DNA was Proteobacteria. However, Actinobacteria, Bacteroidetes, Acidobacteria, Verrucomicrobia, and Planctomycetes were also found within the bacterial community that derived carbon from CH4. In addition, some pyrosequencing reads could not be assigned by the RDP classifier; 12.3 to 22.2% of pyrosequencing reads from the uppermost sediment were assigned to unclassified bacteria, while only about 1.4 to 1.7% of those from the deep sediment were unclassified.
Table 1
| Phylogenetic affiliationa | % (no.) of pyrosequencing readsb from 13C-labeled DNA at: | |||||
|---|---|---|---|---|---|---|
| 4°C | 10°C | 21°C | ||||
| 0-1 cm | 15-20 cm | 0-1 cm | 15-20 cm | 0-1 cm | 15-20 cm | |
| Proteobacteria | ||||||
| Alphaproteobacteria | ||||||
| Methylocystaceae | ||||||
| Methylocystis (type II) | 0.5 (12) | 0 (1) | 1.4 (10) | 1.1 (19) | 2.5 (30) | 0.2 (4) |
| Unclassified Methylocystaceae | —c | — | 0.3 (2) | 0.1 (2) | 0.2 (3) | — |
| Hyphomicrobiaceae, Hyphomicrobiumd | — | — | 2.0 (14) | 0.3 (5) | 4.2 (51) | 0.3 (8) |
| Other Alphaproteobacteria | 6.1 (149) | 11.1 (325) | 6.6 (46) | 10.8 (193) | 13.5 (164) | 15.2 (359) |
| Betaproteobacteria | ||||||
| Methylophilales | ||||||
| Methylophilusd | 13.2 (322) | 20.0 (586) | 4.6 (32) | 26.2 (467) | 0.4 (5) | 9.8 (230) |
| Unclassified Methylophilaceaed | 0.3 (7) | 0.2 (5) | 0.1 (1) | 1.7 (31) | — | 6.6 (156) |
| Burkholderiales, Methylibiumd | 0.8 (19) | 0 (1) | — | — | 0.2 (3) | 0.1 (2) |
| Other Betaproteobacteria | 7.7 (188) | 16.2 (474) | 31.8 (222) | 7.5 (133) | 17.9 (217) | 19.1 (449) |
| Gammaproteobacteria | ||||||
| Methylococcaceae | ||||||
| Methylobacter (type I) | 8.7 (212) | 39.7 (1166) | 1.7 (12) | 5.9 (106) | 0.4 (5) | 21.6 (509) |
| Methylomonas (type I) | 0.4 (10) | — | 4.0 (28) | 0.1 (1) | 5.2 (63) | 0.3 (6) |
| Methylosoma (type I) | 0.2 (4) | — | 5.4 (38) | 25.3 (450) | — | — |
| Unclassified Methylococcaceae (type I) | 0.4 (9) | 0.1 (2) | 0.4 (3) | 0.7 (12) | 0.6 (7) | — |
| Other Gammaproteobacteria | 6.6 (161) | 1.0 (29) | 6.4 (45) | 1.7 (31) | 7.6 (92) | 2.7 (64) |
| Deltaproteobacteria | 9.6 (234) | 1.8 (52) | 7.7 (54) | 0.3 (5) | 10.8 (131) | 0.8 (18) |
| Unclassified Proteobacteria | 0.8 (19) | 0.7 (22) | 1.1 (8) | 0.8 (15) | 2.9 (35) | 0.6 (13) |
| Actinobacteria | 1.4 (35) | 3.3 (96) | 1.6 (11) | 4.9 (88) | 1.2 (14) | 6.4 (150) |
| Bacteroidetes | 4.5 (109) | 0.6 (19) | 3.4 (24) | 3.0 (53) | 3.1 (37) | 5.2 (122) |
| Acidobacteria | 3.5 (86) | 0.1 (4) | 2.9 (20) | 0.4 (8) | 9.1 (110) | 0.5 (11) |
| Verrucomicrobia | 5.7 (138) | 0.1 (4) | 2.3 (16) | 0.5 (9) | 0.8 (10) | 1.7 (41) |
| Planctomycetes | 3.5 (85) | 1.5 (43) | 1.3 (9) | 4.7 (84) | 1.6 (19) | 4.8 (113) |
| Unclassified bacteria | 22.2 (542) | 1.4 (40) | 12.3 (86) | 1.8 (32) | 14.1 (171) | 1.7 (40) |
| Other | 4.1 (99) | 2.2 (65) | 2.6 (18) | 2.1 (38) | 3.6 (44) | 2.5 (60) |
| Total | 100 (2,440) | 100 (2,934) | 100 (699) | 100 (1,782) | 100 (1,211) | 100 (2,355) |
a
“Type I” and “type II” stand for type I and type II methanotrophs.
b
The numbers of pyrosequencing reads were assigned using the RDP classifier (80% confidence threshold).
c
—, no pyrosequencing reads assigned.
d
Methylotroph.
Fig 3
Variation in metabolically active methanotrophs and methylotrophs with temperature.
Methanotrophs are divided into two main taxonomic groups, type I and type II, based on their cell morphology, ultrastructure, phylogeny, and metabolic pathways (27, 44, 61). In our study, the 16S rRNA genes of type I methanotrophs and the pmoA gene were both detected by Q-PCR in ranges of 1.1 × 106 to 1.8 × 107 and 1.2 × 106 to 2.0 × 107 copies μl−1, respectively, in all 13C-labeled DNA samples (Fig. 4). The 16S rRNA genes of type II methanotrophs were detected at 1.8 × 104 and 6.4 × 104 copies μl−1 in the 13C-labeled DNA from sediments at depths of 1 to 3 cm at 10°C and 5 to 10 cm at 21°C, respectively, while they were below the limit of detection of 103 copies reaction−1 in other 13C-labeled DNA. For all the heavy fractions from unlabeled control DNA, the 16S rRNA genes of type I and type II methanotrophs and the pmoA gene were below detection limits.
Fig 4
Pyrosequencing showed that the type I methanotrophs Methylobacter, Methylomonas, Methylosoma, and unclassified Methylococcaceae and the type II methanotroph Methylocystis were all present in the 13C-labeled DNA fractions (Fig. 5). Compared with those of type II methanotrophs, type I methanotroph sequences were more frequently detected in the 13C-labeled DNA pyrosequencing libraries. Methylocystis, the dominant type II methanotroph, was found mainly in the uppermost sediment (depth, 0 to 1 cm), with a relative abundance of 0.5% of pyrosequencing reads for 13C-labeled DNA from the sediment subjected to SIP that was incubated at 4°C, and with higher relative abundances at higher temperatures: 1.4% at 10°C and 2.5% at 21°C. In the deep sediment (depth, 15 to 20 cm), Methylocystis constituted a smaller proportion (0 to 1.1%) of the active bacterial community involved in CH4 oxidation.
Fig 5
Type I methanotrophs were more abundant in the deep sediments (depth, 15 to 20 cm). Methylomonas was dominant in the uppermost sediment incubated at 10°C and 21°C. Methylosoma was found mainly in the sediment incubated at 10°C, while Methylobacter was dominant in the deep sediment and the sediment incubated at 4°C, especially in the deep sediment at 4°C, where it accounted for 39.7% of the total. In addition, we also detected Verrucomicrobia in the 13C-labeled DNA. However, our sequences belonging to the phylum Verrucomicrobia did not have high similarity to the three new Verrucomicrobia isolates identified as CH4 oxidizers (22, 33, 53); rather, they were classified as members of the genus Opitutus, subdivision 3 genera incertae sedis and the genus Prosthecobacter and as unclassified Verrucomicrobiaceae and unclassified Opitutaceae by RDP's Naïve Bayesian Classifier (confidence threshold, 80%).
In addition to methanotrophs, Methylophilus, a known obligate methylotroph that uses methanol as the sole source of carbon and energy, was found to be abundant in 13C-labeled DNA from the deep sediment, particularly at low temperatures. The relative abundance of Methylophilus in the 13C-labeled DNA from the uppermost sediment at 4°C was about 13.2%, but it dropped to 4.6% and 0.4% at 10°C and 21°C, respectively. In the deep sediment, the relative abundance of Methylophilus reached 20 to 26.2% at low temperatures (4°C and 10°C). In addition to Methylophilus, unclassified Methylophilaceae, Methylibium, and Hyphomicrobium (14, 32) were also present in the 13C-labeled DNA. At lower incubation temperatures (4°C), the relative abundance of methylotrophs in the 13C-labeled DNA from the uppermost sediment increased slightly over that at 21°C. We observed a strong correlation (R = 0.82) between the relative abundances of methanotrophs and methylotrophs in pyrosequencing reads of 13C-labeled DNA (Fig. 6).
Fig 6
DISCUSSION
In freshwater sediments, the activity of methanotrophs is largely restricted to a narrow layer at the oxic-anoxic interface (4, 10, 21, 27). In our study, which used a 10% (vol/vol) CH4 concentration in the headspace during incubations, CH4 consumption was observed at all depth intervals tested (down to a sediment depth of 25 cm) and at all three incubation temperatures (4°C, 10°C, and 21°C) (Fig. 2), indicating that the potential for methanotrophy exists down to a sediment depth of at least 25 cm in arctic lakes. Since aerobic methanotrophs cannot be metabolically active in the absence of O2, the methanotrophs from the deeper anoxic sediment layers might be dormant as cysts or exospores in situ (8) and could have become active during the aerobic incubations (57).
The CH4 oxidation potentials in the sediment samples increased with increasing incubation temperature from 4°C to 21°C. A greater increase in the CH4 oxidation potential occurred from 10°C to 21°C than from 4°C to 10°C, indicating that many of the active methanotrophs in the arctic lake sediments might be psychrotolerant (growing at <0 to ≤35°C, with an optimal temperature of ≤25°C) rather than psychrophilic (growing at <0 to ≤20°C, with an optimal temperature of ≤15°C) (50, 58). Similar results were obtained from analyses of the active-layer soils below the surface (top 10 cm) from Ellesmere Island in the Canadian high Arctic, where the active methanotrophs were psychrotolerant rather than psychrophilic (43). In permafrost from Lena Delta, Siberia, the methanotrophic communities in the upper layers were also dominated by psychrotolerant organisms or a mixed community of mesophiles and psychrotolerant organisms, while those close to the permanently frozen ground (30 to 54 cm) were dominated by psychrophiles (41).
Q-PCR of 16S rRNA genes of type I and type II methanotrophs and of pmoA genes, and pyrosequencing of 16S rRNA genes in the heavy fractions (13C-labeled DNA), demonstrated that type I methanotrophs were more abundant and active than type II methanotrophs in the sediments (Table 1; Fig. 4 and 5). Approximately 95.1 to 100% of methanotrophs active at 4°C were assigned to type I methanotrophs. In Lena Delta, Siberia, type I methanotrophs were also more abundant than type II methanotrophs throughout the permafrost active layer of 45 cm or 49 cm according to phospholipid fatty acid (PLFA) analysis (69). In the active-layer soils from Eureka Island in the Canadian high Arctic, Q-PCR of 16S rRNA genes also showed the dominance of type I methanotrophs over type II methanotrophs in the native soil samples, and all methanotrophic 16S rRNA and pmoA gene sequences found in the heavy fractions were related to the type I methanotrophs Methylosarcina and Methylobacter, while no sequence related to type II methanotrophs was identified (43). Denaturing gradient gel electrophoresis (DGGE) of 16S rRNA and pmoA genes showed that the type I methanotrophs Methylobacter and Methylosarcina were dominant in an arctic permafrost active layer of the Lena Delta, Siberia (40). DGGE of the 16S rRNA gene and RNA-SIP also indicated that Methylobacter tundripaludum dominated in soil samples and enrichments from Svalbard (26, 72, 73).
Methylocystis was the only type II methanotroph detected in our sediment samples. The increase in the relative abundance of Methylocystis with an increase in temperature from 4°C to 21°C indicated that high temperatures favored the growth of these type II methanotrophs. Similar results have been obtained for landfill cover soils, where it was found that type I methanotrophs were more dominant at 10°C than at 20°C, while levels of type II methanotrophs were highly elevated only at 20°C (6). In our study, not only did the relative abundances of type I and type II methanotrophs change with temperature, but the composition of the type I methanotrophic community was influenced by temperature. This was most likely due to differences in relative optimum temperatures (2, 27, 54, 62, 65). Microorganisms in permafrost are primarily cold adapted, including psychrophilic and psychrotolerant organisms; very few mesophilic or thermophilic isolates have been identified (25, 41, 64, 67). Based on the relative abundance of methanotrophs in our pyrosequencing reads, we hypothesize that Methylosoma in the sediment was psychrophilic, with an optimum temperature of about 10°C. Methylomonas was likely psychrotolerant or mesophilic, with a decrease in activity and growth at 4°C relative to 21°C. Methylobacter was likely psychrotolerant, as evidenced by substantial growth at 21°C and 4°C.
In addition to methanotrophs, we found that methylotrophs, especially Methylophilus, were active in the acquisition of carbon originally derived from CH4. The relative abundances of methanotrophs, including both type I and type II methanotrophs, correlated significantly with those of methylotrophs, including Methylophilus, unclassified Methylophilaceae, Methylibium, and Hyphomicrobium (Fig. 6) (R = 0.82). Methanol is the first compound produced by methanotrophs during aerobic CH4 oxidation (27). With a high concentration of CH4 and extended incubation times, SIP techniques can lead to significant cross-feeding of 13C into secondary populations (44, 55). Methylotrophs have been widely detected in SIP studies targeting methanotrophic bacteria (11, 26, 42, 43). Cébron et al. (11) suggested that an abundance of methylotrophs such as Methylophilus, Methylovorus, Aminomonas, and Hyphomicrobium in 16S rRNA gene libraries from SIP heavy fractions might result from the incorporation of methanol produced by 13CH4 oxidation. Martineau et al. (43) considered that it was possible that methylotrophs might have acquired carbon by direct utilization of 13CH4 through some previously unknown pathway, because the diversity of methanotrophs is broader than what has generally been reported (15). In our study, methylotrophs were more dominant in 13C-labeled DNA libraries from the deep (15- to 20-cm) sediment, accounting for 20.1 to 27.9% of pyrosequencing reads at low temperatures of 10°C and 4°C, than in those from the uppermost (0- to 1-cm) sediment at 21°C. There may be characteristics of deep sediments or their communities that are associated with the interruption of methanotrophic enzymatic reactions, due to the presence of mutations or inhibitors, such as NaCl, EDTA, and HCOONa, or due to gas composition, leading to an excess of methanol that accumulates extracellularly (16, 23, 24, 37). Methylosoma has also been reported to produce formaldehyde that accumulates extracellularly, which might provide 13C for methylotrophs (48). In addition, because of the in vitro SIP incubations, these methylotrophs could have derived carbon through cross-feeding, in which nonmethanotrophs incorporate 13C into their DNA through the metabolism of by-products, such as 13CO2 or organic matter, derived from methanotrophs (44, 55). It is challenging to determine whether these methylotrophs in 13C-labeled DNA incorporated 13C directly or indirectly from 13CH4. Additional studies are required for understanding of the role of methylotrophs in CH4 cycling in arctic lakes, especially during the period when the lakes are covered with ice.
Despite the enormous potential of SIP techniques for the investigation of microbial community function without the need for cultivation, SIP may distort the original microbial community structure due to in vitro incubation, which can only partially reflect conditions in situ; thus, the data should be interpreted with some caution (13, 44, 55, 56). However, most of the results of this study are consistent with those of the studies in permafrost regions, such as the dominance of type I methanotrophs, including Methylobacter, Methylomonas, and Methylosoma, rather than type II methanotrophs and of psychrotolerant rather than psychrophilic organisms in the arctic lake sediment (40, 41, 43, 69).
Taken together, our findings indicate that CH4 oxidation can occur at low temperatures characteristic of arctic lake sediments, down to at least 4°C. As temperatures increase, the CH4 oxidation potential also increases and is associated with shifts in the composition of bacteria actively growing at the expense of CH4. These shifts are evident within temperature ranges that already occur seasonally in arctic lakes, suggesting that active methanotrophic populations may also change seasonally. As temperatures continue to rise in the Arctic, predominant oxidation rates and active methanotrophic populations will shift. Whether these changes can offset predicted increases in the activity of methanogens is a critical question underlying models of future CH4 flux and resultant climate change.
ACKNOWLEDGMENTS
We thank Ben Gaglioti, Nathan Stewart, Doug Whiteman (Atqasuk, AK), and Monica Heintz for field assistance; Catharine Catranis, John Guido Cable, Heather Slater, Catherine Glover, Robert Burgess, and Mary-Cathrine Leewis for laboratory assistance; and the Alaska Stable Isotope Facility staff, including Tim Howe and Norma Haubenstock. We thank the community in Atqasuk for hosting us. Notably, we thank the Mayor of Atqasuk and the members of the City Council. We thank Thomas Itta, Wanda Kippi, Doug Whiteman, and Kimberly Brent for their input and for allowing us to share our work with them and with high school students at the Meade River School in Atqasuk.
This work was conducted under a Bureau of Land Management permit (FF095556) and North Slope Borough permits (NSB 09-0478 and NSB 10-018). Any use of trade names is only for descriptive purposes and does not imply endorsement by the U.S. Government.
This work was supported by funding from the U.S. Department of Energy National Energy Technology Laboratory (grant DE-NT000565), awarded to Matthew J. Wooller and Mary Beth Leigh.
Supplemental Material
File (aem-aem00853-12-s01.pdf)
- Download
- 285.76 KB
REFERENCES
1.
Anisimov OA. 2007. Potential feedback of thawing permafrost to the global climate system through methane emission. Environ. Res. Lett. 2:045016. doi:.
2.
Bai Y et al. 2006. Phylogenetic diversity of culturable bacteria from alpine permafrost in the Tianshan Mountains, northwestern China. Res. Microbiol. 157:741–751.
3.
Baker GC, Smith JJ, and Cowan DA. 2003. Review and re-analysis of domain-specific 16S primers. J. Microbiol. Methods 55:541–555.
4.
Bender M and Conrad R. 1994. Methane oxidation activity in various soils and freshwater sediments: occurrence, characteristics, vertical profiles and distribution on grain size fractions. J. Geophys. Res. 99:16531–16540.
5.
Berestovskaya YY, Vasil'eva LV, Chestnykh OV, and Zavarzin GA. 2002. Methanotrophs of the psychrophilic microbial community of the Russian Arctic tundra. Mikrobiologiia 71:538–544. (In Russian.)
6.
Börjesson G, Sundh I, and Svensson B. 2004. Microbial oxidation of CH4 at different temperatures in landfill cover soils. FEMS Microbiol. Ecol. 48:305–312.
7.
Bowman JP, McCammon SA, and Skerratt JH. 1997. Methylosphaera hansonii gen. nov., sp. nov., a psychrophilic, group 1 methanotroph from Antarctic marine-salinity, meromictic lakes. Microbiology 143:1451–1459.
8.
Bowman JP, Sly LI, Nichols PD, and Hayward AC. 1993. Revised taxonomy of the methanotrophs: description of Methylobacter gen. nov., emendation of Methylococcus, validation of Methylosinus and Methylocystis species, and a proposal that the family Methylococcaceae includes only the group I methanotrophs. Int. J. Syst. Bacteriol. 43:735–753.
9.
Brosius LS. 2010. Investigating controls over methane production and bubbling from interior Alaskan lakes using stable isotopes and radiocarbon ages. M.S. thesis. University of Alaska, Fairbanks, AK.
10.
Carini S, Bano N, LeCleir G, and Joye SB. 2005. Aerobic methane oxidation and methanotroph community composition during seasonal stratification in Mono Lake, California (USA). Environ. Microbiol. 7:1127–1138.
11.
Cébron A et al. 2007. Identity of active methanotrophs in landfill cover soil as revealed by DNA-stable isotope probing. FEMS Microbiol. Ecol. 62:12–23.
12.
Chanton JP, Powelson DK, Abichou T, Fields D, and Green R. 2008. Effect of temperature and oxidation rate on carbon-isotope fractionation during methane oxidation by landfill cover materials. Environ. Sci. Technol. 42:7818–7823.
13.
Chen Y and Murrell JC. 2010. When metagenomics meets stable-isotope probing: progress and perspectives. Trends Microbiol. 18:157–163.
14.
Chistoserdova L, Kalyuzhnaya MG, and Lidstrom ME. 2009. The expanding world of methylotrophic metabolism. Annu. Rev. Microbiol. 63:477–499.
15.
Conrad R. 2009. The global methane cycle: recent advances in understanding the microbial processes involved. Environ. Microbiol. Rep. 1:285–292.
16.
Corder RE, Johnson ER, Vega JL, Clausen EC, and Gaddy JL. 1986. Biological production of methanol from methane. http://www.anl.gov/PCS/acsfuel/preprint%20archive/Files/33_3_LOS%20ANGELES_09-88_0469.pdf.
17.
Costello AM and Lidstrom ME. 1999. Molecular characterization of functional and phylogenetic genes from natural populations of methanotrophs in lake sediments. Appl. Environ. Microbiol. 65:5066–5074.
18.
Crump BC, Kling GW, Bahr M, and Hobbie JE. 2003. Bacterioplankton community shifts in an arctic lake correlate with seasonal changes in organic matter source. Appl. Environ. Microbiol. 69:2253–2268.
19.
Dedysh SN et al. 2000. Methylocella palustris gen. nov., sp. nov., a new methane-oxidizing acidophilic bacterium from peat bogs, representing a novel subtype of serine-pathway methanotrophs. Int. J. Syst. Evol. Microbiol. 50:955–969.
20.
Dedysh SN, Knief C, and Dunfield RF. 2005. Methylocella species are facultatively methanotrophic. J. Bacteriol. 187:4665–4670.
21.
Duc NT, Crill P, and Bastviken D. 2010. Implications of temperature and sediment characteristics on methane formation and oxidation in lake sediments. Biogeochemistry 100:185–196.
22.
Dunfield PF et al. 2007. Methane oxidation by an extremely acidophilic bacterium of the phylum Verrucomicrobia. Nature 450:879–882.
23.
Frank JJ et al. 1989. On the mechanism of inhibition of methanol dehydrogenase by cyclopropane-derived inhibitors. Eur. J. Biochem. 184:187–195.
24.
Furuto T, Takeguchi M, and Okura I. 1999. Semicontinuous methanol biosynthesis by Methylosinus trichosporium OB3b. J. Mol. Catal. A Chem. 144:257–261.
25.
Gilichinsky DA. 2002. Permafrost, p 2367–2385. In Bitton G (ed), Encyclopedia of environmental microbiology. Wiley, New York, NY.
26.
Graef C, Hestnes AG, Svenning MM, and Frenzel P. 2011. The active methanotrophic community in a wetland from the High Arctic. Environ. Microbiol. Rep. 3:466–472.
27.
Hanson RS and Hanson TE. 1996. Methanotrophic bacteria. Microbiol. Rev. 60:439–471.
28.
He R et al. 17 May 2012. Diversity of active aerobic methanotrophs along depth profiles of arctic and subarctic lake water column and sediments. ISME J. doi:.
29.
Hoehler TM, Borowski WS, Alperin MJ, Rodriguez NM, and Paull CK. 2000. Model, stable isotope, and radiotracer characterization of anaerobic methane oxidation in gas hydrate-bearing sediments of the Blake Ridge. Proc. ODP Sci. Res. 164:79–85.
30.
Holmes AJ, Costello A, Lidstrom ME, and Murrell JC. 1995. Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132:203–208.
31.
Houghton JT et al. 2001. Climate change 2001: the scientific basis. Contribution of Working Group I to the Third Assessment Report of the Intergovernmental Panel on Climate Change. Cambridge University Press, Cambridge, United Kingdom.
32.
Hutchens E, Radajewski S, Dumont MG, McDonald IR, and Murrell JC. 2004. Analysis of methanotrophic bacteria in Movile Cave by stable-isotope probing. Environ. Microbiol. 6:111–120.
33.
Islam T, Jensen S, Reigstad LJ, Larsen O, and Birkeland NK. 2008. Methane oxidation at 55°C and pH 2 by a thermoacidophilic bacterium belonging to the Verrucomicrobia phylum. Proc. Natl. Acad. Sci. U. S. A. 105:300–304.
34.
Jones BM et al. 2011. Modern thermokarst lake dynamics in the continuous permafrost zone, northern Seward Peninsula, Alaska. J. Geophys. Res. 116:G00M03. doi:.
35.
Kightley D, Nedwell DB, and Cooper M. 1995. Capacity for methane oxidation in landfill cover soils measured in laboratory scale soil microcosms. Appl. Environ. Microbiol. 61:592–601.
36.
Lawrence DM and Slater AG. 2005. A projection of severe near-surface permafrost degradation during the 21st century. Geophys. Res. Lett. 32:L24401. doi:.
37.
Lee SG et al. 2004. Optimization of methanol biosynthesis from methane using Methylosinus trichosporium OB3b. Biotechnol. Lett. 26:947–950.
38.
Leigh MB et al. 2007. Biphenyl-utilizing bacteria and their functional genes in a pine root zone contaminated with polychlorinated biphenyls (PCBs). ISME J. 1:134–148.
39.
Lide DR and Frederikse HPR. 1995. CRC handbook of chemistry and physics, 76th ed. CRC Press, Inc., Boca Raton, FL.
40.
Liebner S, Rublack K, Stuehrmann T, and Wagner D. 2009. Diversity of aerobic methanotrophic bacteria in a permafrost active layer soil of the Lena Delta, Siberia. Microb. Ecol. 35:25–35.
41.
Liebner S and Wagner D. 2007. Abundance, distribution and potential activity of methane oxidizing bacteria in permafrost soils from the Lena Delta, Siberia. Environ. Microbiol. 9:107–117.
42.
Lin JL et al. 2004. Molecular diversity of methanotrophs in Transbaikal soda lake sediments and identification of potentially active populations by stable isotope probing. Environ. Microbiol. 6:1049–1060.
43.
Martineau C, Whyte LG, and Greer CW. 2010. Stable isotope probing analysis of the diversity and activity of methanotrophic bacteria in soils from the Canadian High Arctic. Appl. Environ. Microbiol. 76:5773–5784.
44.
McDonald IR, Radajewski S, and Murrell JC. 2005. Stable isotope probing of nucleic acids in methanotrophs and methylotrophs: a review. Org. Geochem. 36:779–787.
45.
Medvedkova KA, Khmelenina VN, Yu A, and Trotsenko YA. 2007. Sucrose as a factor of thermal adaptation of the thermophilic methanotroph Methylocaldum szegediense O-12. Microbiology 76:500–502.
46.
Miller MC, Prentki RT, and Barsdate RJ. 1980. Physics, p 76–178. In Hobbie JE (ed), Limnology of tundra ponds: Barrow, Alaska. Dowden, Hutchinson & Ross, Stroudsburg, PA.
47.
Mohanty SR, Bodelier PLE, and Conrad R. 2007. Effect of temperature on composition of the methanotrophic community in rice field and forest soil. FEMS Microbiol. Ecol. 62:24–31.
48.
Monali R. 2006. Aerobic methanotrophic bacterial communities in sediments of Lake Constance. Ph.D. thesis. University of Constance, Constance, Germany.
49.
Mor S et al. 2006. Induction of enhanced methane oxidation in compost: temperature and moisture response. Waste Manage. 2006:381–388.
50.
Morita RY. 1975. Psychrophilic bacteria. Bacteriol. Rev. 39:144–167.
51.
Omelchenko L, Saveleva ND, Vasileva LV, and Zavarzin GA. 1992. A psychrophilic methanotrophic community from tundra soil. Mikrobiologiia 61:1072–1076.
52.
Pohlman JW, Ruppel C, Hutchinson DR, Downer R, and Coffin RB. 2008. Assessing sulfate reduction and methane cycling in a high salinity pore water system in the northern Gulf of Mexico. Mar. Petrol. Geol. 25:942–951.
53.
Pol A et al. 2007. Methanotrophy below pH 1 by a new Verrucomicrobia species. Nature 450:874–878.
54.
Ponder MA et al. 2005. Characterization of potential stress responses in ancient Siberian permafrost psychroactive bacteria. FEMS Microbiol. Ecol. 53:103–115.
55.
Radajewski S, McDonald IR, and Murrell JC. 2003. Stable-isotope probing of nucleic acids: a window to the function of uncultured microorganisms. Curr. Opin. Biotechnol. 14:296–302.
56.
Radajewski S et al. 2002. Identification of active methylotroph populations in an acidic forest soil by stable-isotope probing. Microbiology 148:2331–2342.
57.
Rothfuss F, Bender M, and Conrad R. 1997. Survival and activity of bacteria in a deep, aged lake sediment (Lake Constance). Microb. Ecol. 33:69–77.
58.
Russell NJ. 1990. Cold adaptation of microorganisms. Philos. Trans. R. Soc. Lond. B Biol. Sci. 326:595–611.
59.
Sazonova TS, Romanovsky VE, Walsh JE, and Sergueev DO. 2004. Permafrost dynamics in the 20th and 21st centuries along the East-Siberian transect. J. Geophys. Res. 109:D01108. doi:.
60.
Schuur EAG et al. 2009. The impact of permafrost thaw on old carbon release and net carbon exchange from tundra. Nature 459:556–559.
61.
Semrau JD, DiSpirito AA, and Yoon S. 2010. Methanotrophs and copper. FEMS Micobiol. Rev. 34:496–531.
62.
Shivaji S et al. 2011. Vertical distribution of bacteria in a lake sediment from Antarctica by culture-independent and culture-dependent approaches. Res. Microbiol. 162:191–203.
63.
Solomon S et al. 2007. Climate change 2007: the physical science basis. Contribution of Working Group I to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change. Cambridge University Press, Cambridge, United Kingdom.
64.
Steven B, Léveillé R, Pollard WH, and Whyte LG. 2006. Microbial ecology and biodiversity in permafrost. Extremophiles 10:259–267.
65.
Steven B, Pollard WH, Greer CW, and Whyte LG. 2008. Microbial diversity and activity through a permafrost/ground ice core profile from the Canadian high Arctic. Environ. Microbiol. 10:3388–3403.
66.
Stoecker K et al. 2006. Cohn's Crenothrix is a filamentous methane oxidizer with an unusual methane monooxygenase. Proc. Natl. Acad. Sci. U. S. A. 103:2363–2367.
67.
Trotsenko YA and Khmelenina VN. 2005. Aerobic methanotrophic bacteria of cold ecosystems. FEMS Microbiol. Ecol. 53:15–26.
68.
Urmann K, Lazzaro A, Gandolfi I, Schroth MH, and Zeyer J. 2009. Response of methanotrophic activity and community structure to temperature changes in a diffusive CH4/O2 counter gradient in an unsaturated porous medium. FEMS Microbiol. Ecol. 69:202–212.
69.
Wagner D, Lipski A, Embacher A, and Gattinger A. 2005. Methane fluxes in extreme permafrost habitats of the Lena Delta: effects of microbial community structure and organic matter quality. Environ. Microbiol. 7:1582–1592.
70.
Walter KM, Smith LC, and Chapin FS III. 2007. Methane bubbling from northern lakes: present and future contributions to the global methane budget. Philos. Transact. A Math. Phys. Eng. Sci. 365:1657–1676.
71.
Walter KM, Zimov SA, Chanton JP, Verbyla D, and Chapin FS III. 2006. Methane bubbling from Siberian thaw lakes as a positive feedback to climate warming. Nature 443:71–75.
72.
Wartiainen I, Hestnes AG, McDonald IR, and Svenning MM. 2006. Methylobacter tundripaludum sp. nov., a methane-oxidizing bacterium from Arctic wetland soil on the Svalbard islands, Norway (78°N). Int. J. Syst. Evol. Microbiol. 56:109–113.
73.
Wartiainen I, Hestnes AG, and Svenning MM. 2003. Methanotrophic diversity in high arctic wetlands on the islands of Svalbard (Norway)—denaturing gradient gel electrophoresis analysis of soil DNA and enrichment cultures. Can. J. Microbiol. 49:602–612.
74.
Wise MG, McArthur JV, and Shimkets LJ. 1999. Methanotroph diversity in landfill soil: isolation of novel type I and type II methanotrophs whose presence was suggested by culture-independent 16S ribosomal DNA analysis. Appl. Environ. Microbiol. 65:4887–4897.
75.
Wooller MJ, Ruppel C, Pohlman JW, Leigh MB, and Heintz M. 2009. Permafrost gas hydrates and climate change: lake-based seep studies on the Alaskan north slope. Fire Ice 9:6–9.
Information & Contributors
Information
Published In
Applied and Environmental Microbiology
Volume 78 • Number 13 • 1 July 2012
Pages: 4715 - 4723
PubMed: 22522690
Copyright
© 2012 American Society for Microbiology.
History
Received: 14 March 2012
Accepted: 9 April 2012
Published online: 7 June 2012
Contributors
Metrics & Citations
Metrics
Note:
- For recently published articles, the TOTAL download count will appear as zero until a new month starts.
- There is a 3- to 4-day delay in article usage, so article usage will not appear immediately after publication.
- Citation counts come from the Crossref Cited by service.
Citations
Citation
2012. Shifts in Identity and Activity of Methanotrophs in Arctic Lake Sediments in Response to Temperature Changes. Appl Environ Microbiol 78:.
If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. For an editable text file, please select Medlars format which will download as a .txt file. Simply select your manager software from the list below and click Download.