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Keywords:

  • Methane monooxygenase gene;
  • Methanotroph;
  • Phylogenetic marker

Abstract

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

The particulate methane monooxygenase gene pmoA, encoding the 27 kDa polypeptide of the membrane-bound particulate methane monooxygenase, was amplified by PCR from DNA isolated from a blanket peat bog and from enrichment cultures established, from the same environment, using methane as sole carbon and energy source. The resulting 525 bp PCR products were cloned and a representative number of clones were sequenced. Phylogenetic analysis of the derived amino acid sequences of the pmoA clones retrieved directly from environmental DNA samples revealed that they form a distinct cluster within representative PmoA sequences from type II methanotrophs and may originate from a novel group of acidophilic methanotrophs. The study also demonstrated the utility of the pmoA gene as a phylogenetic marker for identifying methanotroph-specific DNA sequences in the environment.


1Introduction

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

Methanotrophs are widespread in natural habitats, play an important role in carbon cycling, and have attracted attention because they represent the largest biological sink for methane in aerobic soils [1]. Methane has become one of the most important greenhouse gases, partly because until recently the concentration of methane in the atmosphere has been increasing at a rate of about 1% per year [2]. Wetlands, for example, contribute an estimated 15–20% of the total methane emitted to the atmosphere each year [3]. Functional gene probes targeted at the enzyme soluble methane monooxygenase (sMMO) have been used to identify methanotrophs in blanket bog peat and several other environmental samples [4, 5]. However, the sMMO is not universal to all methanotrophs and is found predominantly in the genera Methylosinus and Methylococcus[6]. Therefore, we examined the use of a second functional gene probe, pmoA encoding the 27 kDa subunit of the particulate methane monooxygenase (pMMO), an enzyme found in all methanotrophs. The genes encoding pMMO polypeptides of 27 and 45 kDa, designated pmoA and pmoB, have recently been cloned from Methylococcus capsulatus (Bath) and are linked on the chromosome of this methanotroph [7]. The predicted amino acid sequences of these genes exhibit a high degree of identity with the corresponding gene products of amoA and amoB which encode key polypeptides of the ammonia monooxygenase (AMO) from the nitrifier Nitrosomonas europaea[8]. Upstream of pmoA and pmoB lies another gene designated pmoC, encoding a putative pMMO polypeptide of around 27 kDa [9]. DNA-DNA hybridisation studies with pmo genes from Methylococcus have revealed that these genes are present in at least two copies in a number of methanotrophs. This gene duplication has also been observed with amo genes in nitrifying bacteria [10]. Sequence data on pmo and amo genes have allowed the design of degenerate PCR primers which will specifically amplify a 525 bp internal DNA fragment of pmoA or amoA from a variety of methanotrophs and nitrifiers. Analysis of pmoA or amoA sequences from representatives of each of the phylogenetic groups of methanotrophs and ammonia oxidising nitrifiers suggests that the particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related enzymes, despite their different physiological roles in these bacteria [11].

The aim of this study was to PCR amplify pmoA sequences directly from DNA samples extracted from blanket peat bog, an environment known to harbour methanotrophs and to correlate the presence of these sequences with similar pmoA sequences obtained from enrichment cultures using the same environmental samples, after phylogenetic analysis, and to demonstrate that the pmoA PCR primers could be used to identify novel and extant methanotrophs which are responsible for methane oxidation in the environment.

2Materials and methods

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

2.1DNA extraction from environmental samples and enrichment cultures

Samples of blanket bog peat were obtained from the Moorhouse National Nature Reserve (grid reference NY 758333) which covers an area of nearly 4000 ha straddling the Pennine Hills in the north of England. Intact core samples of peat (30 cm depth) were obtained using the methods of Hall et al. [12]. The core was carefully extruded from the sample tube and sectioned into 1.0 cm slices between the surface and 10 cm, and into 2.0 cm slices between 10 and 30 cm, with a sharp serrated knife. Total DNA was extracted from peat core sections using methods described previously [5]. This protocol lyses all methanotrophs and methylotrophs in enrichment cultures and in the University of Warwick culture collection (McDonald and Murrell, unpublished). Peat samples consistently yielded high-quality DNA which could be digested with restriction endonucleases and was suitable as a template in PCR amplification experiments.

Methane oxidising bacteria were isolated from blanket bog peat samples in 50 ml enrichment cultures, established in 250 ml conical flasks sealed with rubber Suba seals. The basal medium was the ammonium nitrate mineral salts (ANMS) medium of Whittenbury et al. [13] adjusted to pH 5.8 with phosphate buffer (200 mM potassium phosphate, pH 5.8). Flasks were supplied with methane by removing 50 ml of air from the headspace and injecting 60 ml of a methane-carbon dioxide mixture (95:5). All cultures were incubated at 30°C for 4–6 weeks with shaking at 200 rpm. DNA was extracted from enrichment cultures (final OD540 of 0.1–0.2) using the method described previously [5].

2.2PCR amplification

pmoA genes were amplified from all DNA samples as described previously [11]. Reaction products were checked for size and purity on 1% (w/v) agarose gels and then ligated into the pCR II vector supplied with the TA cloning kit (InVitrogen, San Diego, CA, USA), according to the manufacturer's instructions.

2.3DNA sequencing and analysis

Small-scale preparations of plasmids were done as described previously [5]. DNA sequencing reactions were carried out by cycle sequencing using the ABI PRISM Dye-Terminator Kit (PE Applied Biosystems, Warrington, Cheshire, UK). Primers used for the sequencing reactions were the M13-40 forward and M13 reverse primers (Gibco-BRL, Paisley, UK). Nucleotide and inferred polypeptide sequences were aligned manually with sequences obtained from the GenBank database and dendrograms were constructed using the programs PROTDIST, PROTPARS, DNADIST, DNAPARS, DNAML, FITCH and BOOTSTRAP from the PHYLIP v3.5c package [14]. New pmoA gene sequences have been deposited in GenBank under the accession numbers AF006046 to AF006051.

3Results and discussion

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

PCR primers A189 and A682 were used to amplify pmoA genes from total DNA extracted from peat core samples and total DNA from peat core enrichment cultures. The peat core samples used were from 5–6 cm, 10–12 cm, 18–20 cm and 28–30 cm. Enrichments were made from two peat core sections, 5–6 cm and 10–12 cm, both at pH 5.8 and 4.5 (the blanket bog peat was at pH 3.6). Two clones from each of the eight pmoA clone libraries, prepared using the TA cloning kit (one for each peat core section and enrichment culture studied), were randomly selected and sequenced. The clones selected were from the following sources: clones PD1 and PD2 were from the 5–6 cm peat core section; PD3 and PD4 were from the 10–12 cm peat core section; PD5 and PD6 were from the 18–20 cm peat core section; PD7 and PD8 were from the 28–30 cm peat core section; PE9 and PE10 were from the enrichment established using the 5–6 cm peat core section at pH 5.8; PE11 and PE12 were from the enrichment established using the 5–6 cm peat core section at pH 4.5; PE13 and PE14 were from the enrichment established using the 10–12 cm peat core section at pH 5.8; and PE15 and PE16 were from the enrichment established using the 10–12 cm peat core section at pH 4.5.

The derived PmoA sequences from the 16 environmental pmoA sequences were aligned with the previously published PmoA/AmoA sequences from several extant methanotrophs and nitrifiers [11]. The PmoA sequences obtained from DNA directly extracted from peat contained several identical sequences: clones PD1 and PD8; clones PD2, PD6 and PD7; and clones PD3 and PD5. Seven of the PmoA sequences obtained from DNA extracted from enrichment cultures were identical (clones PE9–PE15) and consequently only one sequence was used in the DNA sequence and amino acid sequence analysis. Therefore only clones PD1, PD2, PD3, and PD4 obtained directly from peat were used in further analyses together with clones PE9 and PE16 obtained from enrichment cultures.

Comparisons of the derived amino acid sequences of the pmoA clones (Table 1) showed that the six environmental clones were closely related to each other, with identity values of 94.0–99.4% and values for similarity of 97.0–100%. The environmental pmoA sequences are quite different from the sequences of (known) nitrifiers and are therefore definitely not related, whereas the clones are 88.5–97.0% identical to PmoA (pmoA) sequences from extant type II methanotrophs.

Table 1.  Similarity/identity matrix comparing derived PmoA/AmoA amino acid sequences of methanotrophs and nitrifiers generated by PCR amplification
 123456789101112131415
  1. Values in the upper triangle of the matrix are ‘percent identity’ and values in the lower triangle, ‘percent similarity’.

1. Clones PD1 and PE9 99.495.898.298.295.195.289.165.561.264.662.446.147.546.1
2. Clone PE16 99.4 95.297.697.694.595.288.565.561.264.662.446.147.646.1
3. Clone PD2 97.697.0 94.094.097.092.788.566.161.864.660.646.147.045.5
4. Clone PD410099.497.6 98.893.394.088.564.860.664.061.245.547.646.1
5. Clone PD3 99.498.897.099.4 93.393.989.165.561.264.661.846.148.246.7
6. Methylocystis sp. strain M 97.697.097.697.697.0 94.586.766.161.862.858.846.147.045.5
7. Methylocystis parvus OBBP 98.297.697.098.297.698.2 88.566.161.863.060.045.146.345.1
8. Methylosinus trichosporium OB3b 97.096.497.697.097.096.495.7 62.459.464.060.643.644.544.2
9. Methylomicrobium album BG8 81.280.682.481.881.881.882.482.4 83.675.267.349.450.649.4
10. Methylomonas methanica S1 80.680.081.881.280.681.281.882.494.5 73.363.048.248.848.8
11. Methylococcus capsulatus (Bath) 80.579.982.381.181.179.979.483.589.790.3 65.953.052.451.8
12. Nitrosococcus oceanus 80.680.080.681.281.278.878.883.083.083.085.447.347.345.143.6
13. Nitrosomonas europaea 67.366.767.367.367.367.966.567.968.970.172.669.1 84.181.8
14. Nitrosospira multiformis 70.770.170.770.770.771.370.170.773.275.078.070.191.5 95.6
15. Nitrosospira sp. Np22 70.970.370.970.970.971.570.170.973.274.478.070.392.199.4 

Phylogenetic analysis of the derived amino acid sequences of the pmoA clones (Fig. 1), using the PROTDIST, PROTPARS, FITCH, and BOOTSTRAP programs of the PHYLIP package [14], showed that three PmoA sequences (PD1, PD3 and PD4) obtained directly by PCR from DNA extracted from a peat core, and two sequences (PE9 and PE16) amplified from methane enrichment cultures form a distinct group (supported by high BOOTSTRAP values) within the type II methanotroph cluster of PmoA sequences. This suggests that they are likely to be PmoA sequences from novel methanotrophs. The sixth PmoA sequence (PD2) also groups within the type II methanotroph PmoA sequences in a branch with Methylocystis sp. strain M PmoA.

image

Figure 1. Phylogenetic analysis of the derived amino acid sequences of pmoA genes from methanotrophs, nitrifiers and environmental DNA samples and enrichments. The dendrogram shows the results from analysis using PROTDIST [14], the BOOTSTRAP values above 50% from 100 replicates are also shown. The bar represents 3% sequence divergence, as determined by measuring the lengths of the horizontal lines connecting any two species.

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Analysis of nucleotide sequences performed with the DNADIST, DNAML, and DNAPARS programs of the PHYLIP package produced dendrograms (data not shown) that were very similar to those derived from the amino acid sequences (Fig. 1). The PmoA sequences from the methane enrichment cultures (PE9 and PE16, also PE10–15) grouped very closely together on the same branch within the type II methanotrophs, suggesting that only a certain type of methanotroph was selected for by the enrichment conditions used. However, an identical PmoA sequence was also amplified from DNA extracted directly from a peat core section (PD1, also PD8) demonstrating that it was possible to identify the same sequence type by both direct extraction and by enrichment. This also shows that the sequence from the peat DNA is from a methanotroph because the identical sequence was isolated from an enrichment culture growing with methane as the sole carbon and energy source, and was therefore from a methanotroph. It should be noted that the enrichment cultures used had been through five subcultures, indicating that the DNA sequences from these cultures were from culturable organisms and not from DNA carried over from the peat core sample used as an inoculum. This suggests that the enrichment conditions used yield methanotrophs representative of the peat environment and that the pmoA sequences retrieved may be from a novel methanotroph that is important in the low pH peat environment.

Sequence analysis has shown that the environmental pmoA sequences (four from DNA extracted from a peat core and two from methane enrichment cultures) originate from a group of organisms with a pmoA sequence related to those of the type II methanotrophs. These sequences, which were the only sequence types to be detected in this study, are quite distinct for this environment. This theory is supported by previous studies of 16S rRNA and mxaF (methanol dehydrogenase) gene sequences from blanket bog peat samples [5, 15]. Novel 16S ribosomal DNA (rDNA) sequences were detected by probing 16S rDNA libraries made from the same sections of the peat core used in this study [5]. These sequences grouped within the 16S rRNA sequences of extant type II methanotrophs and were likely to be from novel acidophilic organisms, since they were isolated from acidic (pH 3.6) blanket bog peat [5]. Novel mxaF sequences were detected using a similar procedure to that used in this study and these sequences grouped with mxaF sequences of extant type II methanotrophs [15]. This further strengthens the theory that these DNA sequences detected by PCR are from novel acidophilic or acid tolerant methanotrophs and also suggests that these organisms may be the dominant methanotrophs in this blanket bog peat, although we cannot exclude the possibility that there may be populations of other extant or novel methanotrophs that may not be amplified in PCR due to their low numbers in this particular environment. However, other work in our laboratory has demonstrated that the primers used will amplify a wide range of methanotrophs and nitrifiers from environmental samples (Starr and Murrell, unpublished), suggesting that the pmoA primers are not biased towards amplifying certain types of sequences over others. We have also demonstrated that pmoA sequences from cultivated methanotroph cells were amplified successfully from environmental samples into which they had been spiked (Enticknap and Murrell, unpublished). Finally, this study has demonstrated the suitability of the pmoA gene as a phylogenetic marker for identifying methanotroph sequences from the environment.

Acknowledgements

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References

This work was supported by the Natural Environment Research Council through its Terrestrial Initiative in Global Environmental Research program (Award GST/02/622), the EU (ERBBIO 4CT960419), and the British Council. We thank Grahame Hall, Institute of Freshwater Ecology, Windermere, UK for peat samples.

References

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results and discussion
  6. Acknowledgements
  7. References
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