SEARCH

SEARCH BY CITATION

Keywords:

  • Soil;
  • Methane oxidising bacterium;
  • Isolation;
  • Natural medium;
  • 16S rRNA hybridisation;
  • ERIC-PCR fingerprinting

Abstract

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

A new method for isolation of methane oxidising bacteria (methanotrophs) is presented. Soil samples from a wetland area and a landfill were plated on polycarbonate membranes, which were incubated in a methane–air atmosphere using a non-sterile soil suspension as the medium. The membrane acted as a permeable growth support. The membrane method resulted in selective growth conditions, which allowed isolation of methane oxidising bacteria. The method resulted in isolation of both type I and type II methanotrophs from natural wetland and landfill soils. The isolates obtained from the landfill were dominated by type II methanotrophs and included several isolates carrying the gene for soluble methane monooxygenase (sMMO). Repetitive element sequence-based PCR fingerprinting documented genotypic diversity at the strain level. The presented method is a promising tool for easy and rapid isolation of different indigenous methanotrophs from an environment of interest.


1Introduction

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

Methane oxidising bacteria (methanotrophs) are ubiquitous and present in freshwater, marine and terrestrial environments where they can represent an important biological sink for methane. Methane sources are anaerobic zones of e.g. sediments, wetland soils or landfills [1–3] and the atmosphere [4–8]. The broad interest for methanotrophs is linked to their role in reducing the emission of methane, as one of the major greenhouse gases, but also linked to the capacity of some methanotrophs to co-oxidise a number of halogenated hydrocarbons [9,10]. The soluble form of methane monooxygenase (sMMO) is of special interest due to its broad substrate specificity [9], and bacteria carrying this gene are possible candidates for bioremediation [10]. The activity of the methanotrophs in soil has been documented [4,8,11,12] and analyses of the methanotrophic bacteria by the use of molecular methods [13–17] and fatty acid profiles [6,18] are well established. Even though these molecular methods can be used on total DNA to describe community structure, there is still a need for environmental isolates to do more comprehensive studies on physiology and functional characteristics of methanotrophs as well as having the opportunity to do polyphasic taxonomy [19]. Direct isolation and enumeration of methanotrophs on artificial solidified media has so far not been successful [20]. Enumeration needs to be done by most probable number (MPN) counting [20] and for isolation initial enrichment cultivation is recommended [21]. Enrichment for, and isolation of, methanotrophs has been done both by use of the standard nitrate mineral salt medium, NMS [21], but also media and incubation parameters more similar to the natural environment during the enrichment step [16,22] or functionally directed [23]. Enrichment cultures may favour dominant or opportunistic bacteria and thereby subsequent loss of a variety of other methanotroph representatives. Furthermore, the enrichment for methanotrophs still supports growth of other bacteria and selectivity thus seems to be limited [20,21].

In this study we present a more direct isolation method, which circumvents the enrichment step. A strong initial selection in favour of methane utilising bacteria was obtained by using a soil substrate membrane system (SSMS), which in addition to isolation, may also have a potential for quantification of these bacteria. Isolates from wetland and landfill soils were further classified as type I or type II by use of molecular methods [16], and the isolates were screened for the presence of sMMO [24]. Enterobacterial Repetitive Intergenic Consensus (ERIC)-PCR fingerprinting finally verified genetic diversity at the strain level [25].

2Materials and methods

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

2.1Soil material

Soils were collected at four locations. Agricultural soils (sandy loams) were collected at the experimental station Højbakkegaard [26] and from a farmer field located at Sørningen 40 km apart. Furthermore, samples were taken from a natural wetland soil located at Roskilde fjord, and from Skellingsted landfill, Western Sealand, Denmark [27,28]. The soil samples were taken from the upper 20 cm and brought to the laboratory and processed on the same day except one soil from Højbakkegaard, which was stored up to 1.5 year at 5°C before use.

2.1.1Incubation of samples on the natural medium membrane system

Three gram of soil (wet weight) for enumeration of colony forming units (CFU) were diluted 1:9 in 0.9% NaCl in MilliQ water, shaken vigorously on a flask shaker for 15 min and left 30 min to allow coarse particles to sediment. The supernatant was serially diluted and plated on a 75 mm polycarbonate microporous cell culture insert having a 0.4 μm pore size (Transwell, Corning Costar, Cambridge, MA, USA) (Fig. 1). During plating of soil suspension the membranes were placed on a Petri dish with NMS agar [21]. This was done to absorb the liquid spread on the membrane. After spreading of the soil dilutions, inserts were immediately transferred to the corresponding Petri dishes containing 8 g of the agricultural soil suspended in 10 ml of sterile MilliQ water so that the polycarbonate membrane was in contact with the soil slurry. The soil substrate was prepared by mixing the total amount of soil and water needed and then distributing it into the separate plates. The dishes were placed in a rack in a gas tight jar of 4 l volume, and a methane–air atmosphere was established by flushing with methane for about 15 s before closing the jar. This resulted in 50–70% of methane in the jars. To verify the ability of methanotrophic growth, filters with Methylomonas methanica (S1) were used as a positive control in some experiments. To evaluate the selectivity of the SSMS incubation, some membranes were also incubated in jars without methane. Agricultural, wetland and landfill soils were all tested as substrates.

image

Figure 1. Schematic overview of the SSMS procedure for membrane growth of methane oxidising bacteria.

Download figure to PowerPoint

The membranes were observed regularly during the SSMS incubation, and jars were aerated and re-flushed with methane every 3–5 days. The incubation was done at 22°C for up to 47 days.

2.1.2Isolation and purification of methanotrophs

From some of the membranes, cells from different colonies were picked by 2 μl capillary tubes and dotted onto new 0.2 μm pore size polycarbonate membranes (Nucleopore Watmann, Kent, UK) which were incubated by floating on new agricultural soil slurry. In a first purification step, colonies from the second membrane growth were serial diluted and filtered onto new 0.2 μm membranes and again incubated on an agricultural soil slurry. To test the colonies for contaminants, membranes were washed by floating on sterile NMS and moved to a rich medium containing 0.5% tryptone, 0.25% yeast extract, 0.1% glucose and 2.0% agar. After incubation for 2 days at 20°C, single pure colonies were selected by microscopy at 25× magnification (Fig. 2). Cells from colonies without any visible traits of contamination were re-streaked and maintained on NMS agar for further characterisation. Cultures were routinely checked for purity by streaking on rich medium, and in case of contamination the purification routines were repeated. Isolates were tested for ability to consume methane. Cell suspensions were washed in NMS, adjusted to OD600=0.2 and 2 ml was added to 120 ml serum flasks with 40 ml NMS and 1% CH4 in headspace and incubated for a few days at room temperature. Methane was measured before and after incubation as described below.

image

Figure 2. Light microscopy picture of pure methanotroph colonies; discrete, well defined spheres, and contaminated colonies; highly irregular, flattened and somewhat amorphous formed (microscopic magnification: 25×).

Download figure to PowerPoint

2.2MPN counts of methanotrophs

Three gram of soil (wet weight) was serial diluted in NMS medium. Sterile filtered NMS medium (400 μl) was added to 3 ml glass vials (Venoject, Terumo, Belgium). Dilutions were made in five to six replicates in 5× dilution steps. The glass vials were closed by silicone coated butyl rubber stoppers, and 0.5 ml methane was added to each vial. Growth was followed visually during 50 days of incubation. At the end of incubation, methane was measured to verify that methane was consumed in vials were growth had occurred. Calculation of cell numbers was done by the method of Klee [29].

2.3Potential methane oxidation rates

Three gram of soil (wet weight) was incubated at 20°C in 38 ml serum flasks sealed with Teflon coated butyl rubber stoppers. The headspace contained air supplemented with 0.12% CH4. Samples taken during incubation were stored in 3 ml venojects (Terumo, Belgium) at −18°C. Methane was analysed on a Varian 3400 GC equipped with an FI detector operated at 160°C and a porapak-Q column. Oven temperature was set to 45°C.

2.4Fluorescent in situ hybridisation

The oligonucleotide probes MG 64 [30] and Mα 450 [31] specific for type I and type II methanotrophs, respectively, were used to verify methanotroph colonies directly on membranes and used to classify the purified isolates by whole cell hybridisation. For hybridisation of colonies on membranes a Bacteria probe, EUB338 [32] and an Archaea specific probe [33] were included as positive and negative control in the protocol. The oligonucleotide probes were fluorescein isothiocyanate (FITC) labelled (DNA technology, Aarhus, Denmark). Hybridisation directly to colonies on membranes followed the procedure of Binnerup et al. [34] after the membranes had been cut into smaller pieces. In the protocol for whole cell hybridisation to purified isolates, pure cultures of M. methanica (S1) and Methylosinus trichosporium (OB3b) were included as positive controls for type I and II methanotrophs, respectively and Escherichia coli strain Mc4100 was used as a negative control. The oligonucleotide probes were Cy3 labelled (Interactiva Biotechnologie, Germany). The hybridisation conditions used to classify the isolates were as described previously [30,31]. Epifluorescence microscopy was performed either with a Leitz DM RB/E microscope (Leica, Wetzlar, Germany) using Filter N2.1 for Cy3 excitation (546 nm) or with a Zeiss Axioplan using filter 09 for FITC exitation.

2.5Molecular characterisation

Cell suspensions were washed in phosphate buffered saline (PBS) (10 mM sodium phosphate; 130 mM NaCl) and resuspended in sterile MilliQ water before used as templates in a PCR reaction. Amplification of the 16S rRNA gene was done with the specific primer pairs MethT1bR–MethT1bF for type I methanotrophs, and MethT2R combined with the bacteria specific primer 27F for type II methanotrophs [16,35]. The PCR was done according to the protocol Wise et al. [16]. For detection of the soluble methane monooxygenase gene (sMMO) the two primer sets 536f and 898r [36] and mmoX1 and mmoX2[37] amplifying the mmoX gene, were used. PCR amplification was done as described [36,37], except for an annealing temperature of 60°C for the primer set 536f and 898r.

The methanotrophic isolates were identified at the strain level by ERIC-PCR fingerprinting [25]. PCR was performed with the following primers: ERIC 1R and ERIC 2 [25]. The oligonucleotides were synthesised by Eurogentec (Seraing, Belgium). The PCR reactions were performed in 25 μl volumes containing 10 mM Tris–HCl (pH 8.8 at 25°C), 50 mM KCl, 1.5 mM MgCl2, 0.1% Triton X-100, 2 μM of each of the two opposing primers ERIC 1R and ERIC 2, 200 μM dNTPs and two units of DynaZyme 500L (Finnzymes Oy, Espoo, Finland). One microlitre of washed cell suspension from pure cultures was used for each PCR amplification. The amplification was performed in a PTC-100 Programmable Thermal Controller (MJ Research Inc., Watertown, MA, USA). The PCR was done according to the protocol for soil bacteria [38]. The PCR products were separated on a 1.5% (w/v) agarose gel (SeaKem LE agarose, FMC BioProducts, ME, USA), stained with 0.4 μg ml−1 ethidium bromide, and visualised by GelDoc system (Bio-Rad, Hercules, CA, USA).

2.6Soil analyses

Soil analyses were done at Chemical Analytical Laboratory, Holt Research Centre, The Norwegian Crop Research Institute according to standard protocols [39,40].

3Results

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

3.1Soil characteristics

Agricultural, wetland and landfill soils were all described and tested for presence of methanotrophs (Table 1). The agricultural soils had a low content of organic carbon and no methane oxidation activity. Inorganic nitrogen was primary present as nitrate, and the highest amounts were found in the stored Højbakkegaard soil. The wetland and landfill soils contained more organic carbon and possessed a potential for methane oxidation in the range of 0.36–2.48 μg C g−1 soil h−1. Presence of methanotrophs was furthermore confirmed by MPN quantification, which gave between 5.4×104 and 4.1×106 methanotrophs g−1 soil.

Table 1.  Soil characteristics and presence of methanotrophs
  1. Bd: below detection limit.

  2. aCH4 oxidation rates were determined at ambient water content. Standard deviations (n=3) are indicated.

  3. bSame batch of soil was used in all experiments. Data range represents the variation between experiments.

  4. cMPN counts after incubation for 50 days in CH4 atmosphere of samples diluted in nitrate minimal salt (NMS) medium.

  5. dNumbers in brackets indicate the upper and lower level of the 95% confidence interval.

  6. eNot determined.

Soil samplesWater content (% (dw))Org-C (% (dw))pHNO3 (μg N g−1 soil (dw))NH4+ (μg N g−1 soil (dw))CH4 oxidationa (μg C g−1 soil (dw) h−1)Number of methanotrophs (MPN g−1 soil (dw)c)
Agricultural soil (sandy loams)       
Højbakkegaard, storedb7.3–9.12.5–2.66.3–6.527–391.0–1.1BdBd
Højbakkegaard, July7.23.27.34.80.9BdBd
Sørninge, July9.44.86.812.02.5BdBd
Wetland soil       
April54.619.57.9eeee
July44.316.87.843.014.00.36±0.045.4×104 (0.5–20.5×104)d
September52.2e7.421.05.11.16±0.061.1×105 (0.2–3.9×105)d
Landfill soil       
April25.86.17.6eeee
July12.28.07.217.011.02.48±0.052.7×105 (0.4–10.6×105)d
September12.4e8.06.31.30.49±0.094.1×106 (0.4–15.0×106)d

3.2Incubation of soil samples on SSMS

Dilutions of agricultural, natural wetland and landfill soil samples were plated directly on polycarbonate membranes. The membranes were carefully transferred to the non-sterile soil slurry (Fig. 1). Only the use of agricultural soil in the soil slurries and incubation in a methane atmosphere supported the formation of a significant number of visible colonies on the membranes (Fig. 3). Incubation of wetland and landfill samples thus resulted in colonies already after a few days of incubation, reaching the maximum number of CFU after about 40 days (Fig. 3). Formation of visible colonies was only seen on membranes with samples having a potential for methane oxidation. Thus between 3×105 and 2×107 CFU g−1 soil (dw) were found in the wetland and landfill samples, in three repeated experiments (Table 2) while no visible colonies were formed on membranes with agricultural soil samples having no capacity for methane oxidation (data not shown). This indicates that the majority of CFU results from growth under conditions of strong selectivity in favour of methane oxidising bacteria, established by using a non-sterile soil slurry as substrate. The different agricultural soils, which were tested as soil substrates, gave no difference in CFU numbers obtained after SSMS incubation of the landfill samples (Table 2). However, some variations in CFU numbers were seen after incubation of the wetland samples (Table 2, July). The highest number of wetland CFU was found on stored Højbakkegaard soil substrate while the lowest number was found on the soil substrate from the fresh Højbakkegaard soil. This difference correlates with the amount of nitrate present in the three agricultural soils (Table 1). Analyses of the agricultural soil substrates after SSMS incubation in fact revealed that all inorganic nitrogen was consumed in the substrates used for wetland sample incubations while inorganic nitrogen still was present in the substrates used for landfill sample incubations (results not shown),

image

Figure 3. Number of colony forming units (CFU) on polycarbonate membranes followed over a period of 47 days of SSMS incubation. Samples collected in September from the wetland location were incubated on SSMS containing slurries of: stored Højbakkegaard soil+methane (•); stored Højbakkegaard soil−methane (◯); fresh wetland soil+methane (▴) and fresh wetland soil−methane (▵).

Download figure to PowerPoint

Table 2.  Numbers of bacteria in wetland and landfill soils determined after SSMS incubation in a methane atmosphere
  1. aCFU appearing after 35–47 days of incubation in a CH4 enriched atmosphere. Standard deviations (n=3) are indicated for July and September samples. In April, deviations are calculated from numbers of CFUs obtained from one replicate sample.

  2. bNot determined.

Soil sample/substrateNumber of bacteria after SSMS incubation (CFU (×106) g−1 soil (dw))a
 Højbakkegaard, stored soilHøjbakkegaard, fresh soilSørninge, fresh soil
Wetland   
April1.50±0.20bb
July1.30±0.300.30±0.040.60±0.07
September19.70±2.60bb
Landfill   
April7.23±0.31bb
July4.02±0.854.40±1.103.40±1.28
September0.90±0.20bb

To verify that colonies on the membranes de facto were methanotrophs, oligonucleotide probes specific for Bacteria, type I methanotrophs and type II methanotrophs were used for colony hybridisation after 30 days SSMS incubation of samples from July. However, none of the visible colonies fluoresced when hybridised to the Bacteria (EUB338) or the methanotroph specific probes neither did cell mass from the colonies (results not shown). The lack of signal could be due to restricted probe accessibility or low ribosome content.

3.3Characterisation of methanotroph isolates

In one of the experiments (April) presence of methanotrophs in the colonies was verified by picking 24 colonies in total from the membranes after SSMS incubation of the wetland and landfill samples. The isolates obtained after purification included rods, cocci, curved and straight cells. Most of the isolates were opaque or weak brownish except MSB4 and MSB7, which had a clear red-orange pigmentation. All the isolates were found to consume methane but none of them were able to grow above 40°C (data not shown).

Based on PCR with 16S rDNA targeted primers with group specificity for type I or type II methanotrophs and whole cell hybridisation with fluorescently labelled 16S rRNA targeted oligonucleotide probes, 15 of the landfill isolates were identified as type II methanotrophs and one (MSB25) as a type I methanotroph (Fig. 4). From the wetland soil three out of the eight isolates were type II methanotrophs (Fig. 4). One methane oxidising isolate from the wetland (MSB5), could not be identified as a type I or type II methanotroph by the group specific PCR or the oligonucleotide probes. The ERIC-PCR fingerprint method was successfully applied to all the isolates and the banding patterns revealed a genetic diversity at the strain level (Fig. 4). Only two of the isolates, MSB4 and MSB7 (type I), were found to be identical by morphology and ERIC-PCR fingerprinting (data not shown). The mmoX gene coding for the α-subunit of the hydroxylase component of the sMMO was found by PCR amplification in four strains from the wetland and 10 strains from the landfill soil (Fig. 4). All the mmoX carrying strains except one (MSB8) correlated with the type II classification. However, five of the type II landfill strains did not possess the mmoX gene.

image

Figure 4. ERIC-PCR fingerprint patterns of methanotrophic strains isolated from a wetland soil (MSB1–8) and a landfill soil (MSB11–26). Classification of the strains is based on PCR with 16S rDNA targeted primers with group specificity for type I and type II methanotrophs, whole cell hybridisation with group specific 16S rRNA targeted probes (MG 64 and Mα 450) and detection of the soluble methane monooxygenase gene (sMMO). pGEM; DNA molecular weight markers (Promega).

Download figure to PowerPoint

4Discussion

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

Isolation of indigenous methanotrophs can be rather problematic due to their slow growth rates and growth of other non-methane utilising bacteria during cultivation [20,41,42]. Furthermore, the majority of them do not express colony morphology or pigmentation suitable for selection. Use of the new approach presented in this paper, including the soil substrate membrane system (SSMS), seems to circumvent these problems. The method gave selective growth of methane oxidising bacteria resulting in distinct visible colonies on a membrane surface within a few weeks of incubation. In the absence of methane or plating of a soil sample without methane oxidation potential, none or very few colonies were formed. The initial selection in favour of methane consuming bacteria and the separation of cells on the membrane surface are essential components of this method. In a traditional liquid enrichment culture the cells will be in contact with each other and substrates from the active cells are distributed in the liquid to support growth of other bacteria than the methanotrophs [21]. The enrichment of a simple ecosystem is difficult to avoid. Our hypothesis is that microorganisms present in the non-sterile substrate soil used during the SSMS incubation, act as a buffer consuming the metabolites produced by the methanotrophs during growth and colony formation. This prevents other bacteria than the methanotrophs from forming visible colonies on the membranes. Even though all the colonies selected for further characterisation contained methane oxidising bacteria, most of them still harboured other bacteria within the colony and further purification was essential. One of the first purification steps included a new SSMS incubation of serial diluted colony suspensions followed by a few days incubation of the membrane on a rich agar medium. This was a successful strategy to identify methanotroph colonies containing other heterotrophic microorganisms, which enabled us to isolate purified methanotroph colonies within a short time.

In this study agricultural soils without methane oxidation potential were found most suitable as substrate for the SSMS incubation. Soil with high methanotroph activity (wetland and landfill soils) did not support formation of colonies on the membranes when used as a SSMS substrate. We hypothesise that colony formation on the membranes is prevented by an early nutrient depletion during the incubation as a result of high initial methanotroph activity and growth in the SSMS substrate itself. However, all the agricultural soils did support formation of colonies, although some differences were seen when wetland soil was incubated on different agricultural soils. Combination of a high number of colonies on the membranes used for quantification of wetland CFU (>100) and a low concentration of nitrate in some of the agricultural soils may be the reason for nitrogen depletion and variation in CFU numbers after SSMS incubation of the wetland samples collected in July. This result combined with the fact that ammonium may also prevent growth of methanotrophs [43] stress the importance of analysing the soils used as substrate components for their nutritional status.

The use of native media has been reported previously to support growth of a high percentage of indigenous bacteria [44,45] and the recent study by Kaeberlein et al. [46] has shown that bacteria apparently unculturable on artificial media can proliferate on a ‘simulated natural environment’ medium. In addition to the potential for establishing highly selective growth conditions, the SSMS incubation may give new possibilities for isolation of indigenous bacteria normally difficult to cultivate and isolate on the more frequently used media.

Here, cells from 24 colonies appearing on the membranes after SSMS incubation were purified and the isolates were fingerprinted by ERIC-PCR. The fingerprints show a genotypic diversity including 23 different strains, which stress the avoidance of the enrichment step to prevent dominance of a few genotypes as found in other studies [16]. ERIC-PCR fingerprinting has been applied with success on different soil bacteria and found useful both for diversity studies at low taxonomic levels and for identification of strains [38,47,48]. This study shows the usefulness of the method on environmental isolates of methanotrophs.

The classification based on specific 16S rDNA PCR and whole cell hybridisation showed different methanotroph compositions in the two soils. Type II methanotrophs dominated in landfill soil which is in accordance with other studies [16,49] while a mixture of type I and type II methanotrophs was present in the wetland soil. The methanotrophs containing sMMO have a broad substrate specificity and are of special interest in degradation of halogenated hydrocarbons in landfill soils [10]. In this study we were able to identify a subunit of the gene for this enzyme with two different primer sets in three type II strains from the wetland soil and in 10 strains from the landfill soil. The landfill soil used in our study has the capacity to degrade different halogenated ethylenes [50], which seems to correlate with the presence of a methanotroph community including different strains carrying the sMMO gene.

All colonies subjected to further characterisation were identified as methanotrophs by growth in methane–air atmosphere and molecular methods. Even though colonies were only seen on the membranes when methane was present during the incubations and when samples with significant methanotroph activity were filtered onto the membranes, attempts to identify methanotroph colonies directly on the membranes failed. Whole cell fluorescent hybridisation did not result in a probe specific signal from cells in the large colonies, probably due to low ribosome content as a result of a starvation response [51].

Isolation, characterisation and quantification of methanotrophic bacteria are of high relevance because of the special emphasis on their role in methane emission to the atmosphere [52], in bioremediation [53] and in industry related to single cell protein production based on natural gas (http://www.norferm.no). The presented method based on direct isolation on membranes without enrichments, is an obvious advantage in terms of time, effort and number of isolates compared to enrichments in liquid cultures and isolation on agar medium.

Acknowledgements

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

We thank Margit Møller Fernqvist for technical assistance. Dr Mette Christophersen and Dr Charlotte Scheutz are thanked for help with soil sampling from Skellingsted Landfill and Søren Christensen for help with the methane measurements. The project was supported by the Nordic Council of Ministers (project 651044-10224).

References

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References
  • [1]
    Hanson, R.S., Hanson, T.E. (1996) Methanotrophic bacteria. Microbiol. Rev. 60, 439471.
  • [2]
    Conrad, R. (1996) Soil microorganisms as controllers of atmospheric trace gases (H2, CO, CH4, OCS, N2O and NO). Microbiol. Rev. 60, 609640.
  • [3]
    Reeburgh, W.S., Whalen, S.C. and Alperin, M.J. (1993) The role of methylotrophy in the global methane budget, In: Microbial Growth on C1 Compounds (Murrell, J.C. and Kelly, D.P., Eds.), pp. 1–14. Intercept Ltd., Andover.
  • [4]
    Bull, I.D., Parekh, N.R., Hall, G.H., Ineson, P., Evershed, R.P. (2000) Detection and classification of atmospheric methane oxidising bacteria in soil. Nature 405, 175178.
  • [5]
    Conrad, R. (1997) Production and consumption of methane in the terrestrial biosphere. In: Biogenic Volatile Organic Carbon Compounds in the Atmosphere (Helas, G., Slanina, J. and Steinbrecher, R., Eds.), pp. 27–44. SBP Academic Publishers, Amsterdam.
  • [6]
    Holmes, A.J., Roslev, P., McDonald, I.R., Iversen, N., Henriksen, K., Murrell, J.C. (1999) Characterization of methanotrophic bacterial populations in soils showing atmospheric methane uptake. Appl. Environ. Microbiol. 65, 33123318.
  • [7]
    Roslev, P., Iversen, N. (1997) Oxidation and assimilation of atmospheric methane by soil methane oxidisers. Appl. Environ. Microbiol. 63, 874880.
  • [8]
    Roslev, P., Iversen, N. (1999) Radioactive fingerprinting of microorganisms that oxidize atmospheric methane in different soils. Appl. Environ. Microbiol. 65, 40644070.
  • [9]
    Burrows, K.J., Cornish, A., Scott, D., Higgins, I.J. (1984) Substrate specificities of the soluble and particulate methane monooxygenases of Methylosinus trichosporium OB3b. J. Gen. Microbiol. 5, 335342.
  • [10]
    Oldenhuis, R., Vink, R.L.J.M., Janssen, D., Witholt, B. (1989) Degradation of chlorinated aliphatic hydrocarbons by Methylosinus trichlosporium OB3b expressing soluble methane monooxygenase. Appl. Environ. Microbiol. 55, 28192826.
  • [11]
    McDonald, I.R., Hall, G.H., Pickup, R.W., Murrell, J.C. (1996) Methane oxidation potential and preliminary analysis of methanotrophs in blanket bog peat using molecular ecology techniques. FEMS Microbiol. Ecol. 21, 197211.
  • [12]
    Börjesson, G., Sundh, I., Tunlid, A., Frostegård, Å., Svensson, B.H. (1998) Microbial oxidation of CH4 at high partial pressure in an organic landfill cover soil under different moisture regimes. FEMS Microbiol. Ecol. 26, 207217.
  • [13]
    Henckel, T., Roslev, P., Conrad, R. (2000) Effects of O2 and CH4 on presence and activity of the indigenous methanotrophic community in rice field soil. Environ. Microbiol. 2, 666679.
  • [14]
    Holmes, A.J., Owens, N.J.P., Murrell, J.C. (1996) Molecular analysis of enrichment cultures of marine methane oxidising bacteria. J. Exp. Mar. Biol. Ecol. 203, 2738.
  • [15]
    McDonald, I.R., Kenna, E.M., Murrell, J.C. (1995) Detection of methanotrophic bacteria in environmental samples with the PCR. Appl. Environ. Microbiol. 61, 116121.
  • [16]
    Wise, M.G., McArthur, J.V., Shimkets, L.J. (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, 48874897.
  • [17]
    Murrell, J.C., McDonald, I.R., Bourne, D.G. (1998) Molecular methods for the study of methanotroph ecology. FEMS Microbiol. Ecol. 27, 103114.
  • [18]
    Guckert, J.B., Ringleberg, D.B., White, C.C., Hanson, R.S., Bratina, B.J. (1991) Membrane fatty acids as phenotypic markers for the polyphasic approach to taxonomy of methylotrophs within the Proteobacteria. J. Gen. Microbiol. 137, 26312641.
  • [19]
    Vandamme, P., Pot, B., Gillis, M., De Vos, P., Kersters, K., Swings, J. (1996) Polyphasic taxonomy, a consensus approach to bacterial systematics. Microbiol. Rev. 60, 407438.
  • [20]
    Escoffier, S., Le Mer, J., Roger, P.A. (1997) Enumeration of methanotrophic bacteria in ricefield soils by plating and MPN techniques: a critical approach. Eur. J. Soil Biol. 33, 4151.
  • [21]
    Whittenbury, R., Phillips, K.C., Wilkinson, J.F. (1970) Enrichment, isolation and some properties of methane-utilizing bacteria. J. Gen. Microbiol. 61, 205218.
  • [22]
    Dedysh, S.N., Panikov, N.S., Tiedje, J.A. (1998) Acidophilic methanotrophic communities from Sphagnum peat bogs. Appl. Environ. Microbiol. 64, 922929.
  • [23]
    Bodrossy, L., Murrell, J.C., Dalton, H., Kalman, M., Puskas, L.G., Kovacs, K. (1995) Heat-tolerant methanotrophic bacteria from the hot water effluent of a natural gas field. Appl. Environ. Microbiol. 61, 35493555.
  • [24]
    McDonald, I.R., Uchiyama, H., Kambe, S., Yagi, O., Murrell, J.C. (1997) The soluble methane monooxygenase gene cluster of the trichlorethylene-degrading methanotroph Methylocystis sp. Strain M. Appl. Environ. Microbiol. 63, 18981904.
  • [25]
    Versalovic, J., Koeuth, T., Lupski, J.R. (1991) Distribution of repetitive DNA sequences in eubacteria and applications to fingerprinting of bacterial genomes. Nucleic Acids Res. 19, 68236831.
  • [26]
    Højberg, O., Binnerup, S.J., Sørensen, J. (1996) Potential rates of ammonium oxidation, nitrite oxidation, nitrate reduction and denitrification in the young barley rhizosphere. Soil Biol. Biochem. 28, 4754.
  • [27]
    Kjeldsen, P., Fischer, E.V. (1995) Landfill gas migration-field investigations at Skellingsted Landfill, Denmark. Waste Manag. Res. 13, 467484.
  • [28]
    Christophersen, M., Linderød, L., Jensen, P.E., Kjeldsen, P. (2000) Methane oxidation at low temperatures in soil exposed to landfill gas. J. Environ. Qual. 29, 19891997.
  • [29]
    Klee, A.J. (1993) A computer-program for the determination of most probable number and its confidence-limits was used. J. Microbiol. Methods 18, 9198.
  • [30]
    Bourne, D.G., Holmes, A., Iversen, N., Murrell, J.C. (2000) Fluorescent oligonucleotide rDNA probes for specific detection of methane oxidising bacteria. FEMS Microbiol. Ecol. 31, 2938.
  • [31]
    Eller, G., Stubner, S., Frenzel, P. (2001) Group-specific 16S rRNA targeted probes for the detection of type I and type II methanotrophs by fluorescence in situ hybridization. FEMS Microbiol. Lett. 198, 9197.
  • [32]
    Amann, R.I., Krumholz, L., Stahl, D.A. (1990) Fluorescent-ologonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J. Bacteriol. 172, 762770.
  • [33]
    Stahl, D.A. and Amann, R.I. (1991) Development and application of nucleic acid probes in bacterial systematics. In: Nucleic Acid Techniques in Bacterial Systematics (Stackebrandt, E. and Goodfellow, M., Eds.), pp. 205–248. Wiley, Chichester.
  • [34]
    Binnerup, S.J., Bloem, J., Hansen, B.M., Veninga, M., Wolters, W., Hansen, M. (2001) Ribosomal RNA content in microcolony forming soil bacteria measured by quantitative 16S rRNA hybridization and image analysis. FEMS Microbiol. Ecol. 37, 231237.
  • [35]
    Lane, D.J. (1991) 16S/23S rRNA sequencing. In: Nucleic Acid Techniques in Bacterial Systematics (Stackebrandt, E. and Goodfellow, M., Eds.), pp. 115–175. Wiley, Chichester.
  • [36]
    Fuse, H., Misaki, O., Takimura, O., Murkami, K., Inoue, H., Yamaoka, Y., Oclarit, J.M., Omori, T. (1998) Oxidation of trichloroethylene and dimethyl sulfide by a marine Methylomicrobium strain containing soluble methane monooxygenase. Biosci. Biotechnol. Biochem. 62, 19251931.
  • [37]
    Miguez, C.B., Bourque, D., Sealy, J.A., Greer, C.W., Groleau, D. (1997) Detection and isolation of methanotrophic bacteria possessing soluble methane monooxygenase (sMMO) genes using the polymerase chain reaction (PCR). Microbiol. Ecol. 33, 2131.
  • [38]
    de Bruijn, F.J. (1992) Use of repetitive (Repetitive Extragenic Palindromic and Enterobacteria Repetitive Intergenic Consensus) sequences and the polymerase chain reaction to fingerprint the genomes of Rhizobium meliloti isolates and other soil bacteria. Appl. Environ. Microbiol. 58, 21802187.
  • [39]
    R.K. Schofiled, A.W. Taylor (1955) The measurement of soil pH. Soil Sci. Soc. Am. Proc. 19
  • [40]
    J.M. Bremner (1965) Inorganic forms of nitrogen. Methods of soil analysis. Agronomy 11 9
  • [41]
    Whittenbury, R. and Dalton, H. (1981) The methylotrophic bacteria. In: The Prokaryotes (Starr, M.P., Stolp, H., Truper, G., Balows, A. and Schlegel, H.G., Eds.), pp. 894–902. Springer, Berlin.
  • [42]
    Hanson, R.S., Netrusov, A.I. and Tsuji, K. (1992) The obligate methanotrophic bacteria Methylococcus, Methylomonas and Methylosinus and related bacteria. In: The Prokaryotes (Balows, A., Truper, G., Dworkin, M., Harder, W. and Schleifer, K.H., Eds.), pp. 2350–2365. Springer, New York.
  • [43]
    Goulding, K.W.T., Willison, T.W., Webster, C.P., Powlson, D.S. (1996) Methane fluxes in aerobic soils. Environ. Monit. Assess. 42, 175187.
  • [44]
    Olsen, R.A., Bakken, L.R. (1987) Viability of soil bacteria: optimization of plate-counting technique and comparison between total counts and plate counts within different size groups. FEMS Microbiol. Ecol. 13, 5974.
  • [45]
    Winding, A., Binnerup, S.J., Sørensen, J. (1994) Viability of indigenous soil bacteria assayed by respiratory activity and growth. Appl. Environ. Microbiol. 60, 28692875.
  • [46]
    Kaeberlein, T., Lewis, K., Epstein, S.S. (2002) Isolating ‘uncultivable’ microorganisms in pure culture in a simulated natural environment. Science 296, 11271129.
  • [47]
    Louws, F.J., Fulbright, D.W., Stephens, C.H., de Bruijn, F. (1994) Specific genomic fingerprints of phytopathogenic Xanthomonas and Pseudomonas pathovars and strains generated with repetitive sequences and PCR. Appl. Environ. Microbiol. 60, 22862295.
  • [48]
    Svenning, M.M., Gudmundsson, J., Fagerli, I.L., Leinonen, P. (2001) Competition for nodule occupancy between introduced strains of Rhizobium leguminosarum biovar trifolii and its influence on plant production. Ann. Bot. 88, 781787.
  • [49]
    Mandernack, K.W., Kinney, C.A., Coleman, D., Huang, Y.S., Freeman, K.H., Bogner, J. (2000) The biogeochemical controls of N2O production and emission in landfill cover soils: the role of methanotrophs in the nitrogen cycle. Environ. Microbiol. 2, 298309.
  • [50]
    Scheutz, C. and Kjeldsen, P. (2000) Methane oxidation and degradation of halogenated organic compounds in landfill gas affected soil. In: Proceedings Intercontinental Landfill Research Symposium, Luleå University of Technology, Luleå, Sweden.
  • [51]
    Oda, Y., Slagman, S.-J., Meijer, W.G., Forney, L.J., Gottschal, J.C. (2000) Influence of growth rate and starvation on fluorescent in situ hybridization of Rhodopseudomonas palustris. FEMS Microbiol. Ecol. 32, 205213.
  • [52]
    Topp, E. and Hanson, R.S. (1991) Metabolism of radioactively important trace gases by methane-oxidising bacteria. In: Microbial Production and Consumption of Greenhouse Gases: Methane, Nitrogen Oxides, and Halomethanes (Rogers, J.E. and Whitman, W.B., Eds.), pp. 71–90. American Society of Microbiology, Washington, DC.
  • [53]
    Sullivan, J.P., Dickinson, D., Chase, H.A. (1998) Methanotrophs, Methylosinus trichosporium OB3b, sMMO, and their application to bioremediation. Crit. Rev. Microbiol. 24, 335373.