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Applied and Environmental Microbiology, May 2000, p. 1801-1808, Vol. 66, No. 5
Max-Planck-Institut für terrestrische
Mikrobiologie, Karl-von-Frisch-Strasse, 35043 Marburg, Germany
Received 1 October 1999/Accepted 7 January 2000
Forest and other upland soils are important sinks for atmospheric
CH4, consuming 20 to 60 Tg of CH4 per year.
Consumption of atmospheric CH4 by soil is a microbiological
process. However, little is known about the methanotrophic bacterial
community in forest soils. We measured vertical profiles of atmospheric
CH4 oxidation rates in a German forest soil and
characterized the methanotrophic populations by PCR and denaturing
gradient gel electrophoresis (DGGE) with primer sets targeting the
pmoA gene, coding for the The atmospheric concentration of
CH4, one of the most important greenhouse gases, has
increased dramatically over the past 200 years. About 80 to 90% of
atmospheric CH4 is of biogenic origin (20). The
major sink is the chemical destruction by OH· and Cl· radicals in
the troposphere and stratosphere, respectively (9, 10).
However, the capacity of these atmospheric sinks may decline, since the
rising concentrations of other trace gases emitted by anthropogenic
activity result in a reduction of OH· radicals in the troposphere
(27).
The only biological sink for CH4 is oxidation in soil.
Atmospheric CH4 is consumed in forest, agricultural, and
other upland soils. CH4 consumption in these soils is
caused by methane-oxidizing bacteria. However, the identity of these
methanotrophs is still unknown. The apparent half-saturation constant
(Km) for oxidation of atmospheric
CH4 (approximately 1.8 parts per million by volume [ppmv]) in upland soils ranges from 0.8 to 280 nM (6, 7, 13,
16). However, the Km of the common type I
or II methanotrophs (0.8 to 66 µM), which are available in culture
collections, is 1 to 3 orders of magnitude higher, and these common
methanotrophs are not able to survive for a prolonged period using only
atmospheric CH4 (14, 17, 37). Recently, however,
a type II methanotroph was isolated from a humisol that was able to
adapt to nanomolar Km values, close to those
measured in upland soils (14; P. Dunfield, personal
communication). The existence of this isolate brought into question the
hypothesis of Bender and Conrad (6) that besides the common
low-affinity methanotrophs (micromolar Km), unknown high-affinity methanotrophs (nanomolar
Km) exist and that only the latter are
responsible for atmospheric CH4 oxidation.
Common methanotrophs have a neutral pH optimum (18).
However, most forest soils are slightly acidic, around pH 5 and lower. Bacteria extracted from forest soils showed a methanotrophy pH optimum
of 5.8, indicating that unknown acidophilic methanotrophs may be
responsible for CH4 oxidation in forest soils
(5). Indeed, an acidophilic methanotroph, belonging to the
However, so far little is known about the methanotrophic community in
forest soil. A recent analysis of pmoA gene libraries from
forest soils has demonstrated the existence of a new group of
methanotrophs in various forest soils (23). These soils all exhibited uptake of atmospheric CH4.
Forest soils have been extensively studied with respect to
CH4 oxidation kinetics and zonation and inhibition of
CH4 oxidation because of their important function as major
sinks in the global CH4 budget. The highest CH4
oxidation activity in forest soils was usually measured in subsurface
soil layers (1, 29, 33, 38). This localization of
methanotrophs in deeper soil layers was attributed to inhibition of
methanotrophs by ammonium or terpenes that are released or produced in
the organic surface layers of the forest soil (2-4, 16, 26,
36). However, the zonation of CH4 oxidation in
relation to the involved methanotrophic community has not yet been studied.
Therefore, we measured vertical profiles of the uptake of atmospheric
CH4 in forest soil cores in winter and summer and
characterized the bacterial and involved methanotrophic populations by
PCR and denaturing gradient gel electrophoresis (DGGE) using a
universal small-subunit rRNA gene (SSU rDNA) primer set and a primer
set targeting the pmoA gene.
Sampling site and soil characteristics.
The sampling site
was located on a slope in a deciduous forest near Marburg, Germany (N
51°00.000', E 09° 50.625'), consisting of mainly beech (Fagus
sylvatica) and oak (Quercus robur). The soil type was a
cambisol with a Ah (2 to 6 cm), Bv (6 to 28 cm), and C (sandstone) horizon. The soil originated on sandstone and was a loamy sand. The pHH2O values of the
organic Ah horizon and the mineral subsoil were pH 3.8 and
pH 4.3, respectively. Soil cores were collected in January (winter) and
July (summer) of 1999.
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Molecular Analyses of Novel Methanotrophic
Communities in Forest Soil That Oxidize Atmospheric Methane
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
subunit of the particulate
methane monooxygenase, and the small-subunit rRNA gene (SSU rDNA) of
all life. The forest soil was a sink for atmospheric CH4 in
situ and in vitro at all times. In winter, atmospheric CH4
was oxidized in a well-defined subsurface soil layer (6 to 14 cm deep),
whereas in summer, the complete soil core was active (0 cm to 26 cm
deep). The content of total extractable DNA was about 10-fold higher in
summer than in winter. It decreased with soil depth (0 to 28 cm deep)
from about 40 to 1 µg DNA per g (dry weight) of soil. The PCR product concentration of SSU rDNA of all life was constant both in winter and
in summer. However, the PCR product concentration of pmoA changed with depth and season. pmoA was detected only in
soil layers with active CH4 oxidation, i.e., 6 to 16 cm
deep in winter and throughout the soil core in summer. The same
methanotrophic populations were present in winter and summer. Layers
with high CH4 consumption rates also exhibited more bands
of pmoA in DGGE, indicating that high CH4
oxidation activity was positively correlated with the number of
methanotrophic populations present. The pmoA sequences
derived from excised DGGE bands were only distantly related to those of
known methanotrophs, indicating the existence of unknown
methanotrophs involved in atmospheric CH4 consumption.
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
proteobacteria and closely related to the nonmethanotroph
Bejerinckia but only distantly related to the common type II
methanotrophs, was recently isolated from an acidic blanket peat bog in
Russia (11, 12).
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
1 (n = 5),
respectively (35). The gravimetric water content was determined for each 2-cm vertical soil section and expressed as a
percentage of WHC.
Vertical CH4 concentration profiles. Gas samples were taken by pushing a PEEK capillary (diameter, 0.35 mm; Sykam, Gilchingen, Germany) attached to a stainless steel rod into the soil at 1-cm intervals. Prior to sampling, the tube was flushed by extracting 0.1 ml of gas. Gas samples (1 ml) were collected in gastight syringes and analyzed by gas chromatography with a GC-8A (Shimadzu, Japan) equipped with a flame ionization detector and a stainless steel column (length, 2 m; diameter, 1/8 inch) filled with Poropak Q (80/110 mesh).
Collection of soil cores and in vitro CH4 oxidation rate. Soil cores were taken with Plexiglas corers (diameter, 6 cm) pushed into the forest floor. The columns were extracted from the ground and closed with silicone stoppers. In the laboratory, the cores were gently pushed upward and cut into 2-cm sections. The sectioned soil layers were immediately transferred into 150-ml serum bottles, which were closed with latex stoppers.
Gas samples were repeatedly taken over time with gastight pressure-lock syringes (A-2 series; Dynatech, Baton Rouge, La.) and analyzed by gas chromatography. Apparent first-order CH4 oxidation rate constants were calculated (Excel 7.0; Microsoft) from the exponential decrease of CH4 with time and converted to CH4 oxidation rates by multiplication with the atmospheric CH4 mixing ratio (1.7 ppmv).DNA extraction.
DNA extraction from forest soil was modified
from the method of Moré et al. (31). Approximately
0.5 g (fresh weight) of soil was taken from each forest soil layer
(2 cm thick) and transferred into 2-ml screw cap tubes. Approximately
1 g of sterilized (170°C for 4 h) zirconia/silica beads
(diameter, 0.1 mm; Biospec Products Inc., Bartlesville, Okla.), 800 µl of 120 mM sodium phosphate buffer (pH 8), and 260 µl of sodium
dodecyl sulfate solution (10% sodium dodecyl sulfate, 0.5 M Tris-HCl
[pH 8.0], 0.1 M NaCl) were added, and the soil was resuspended
homogeneously by vortexing. The cells were lysed for 45 s by
shaking in a cell disruptor (FP120 FastPrep; Savant Instruments Inc.,
Farmingdale, N.Y.) at a setting of 6.5 m s1. After
centrifugation (3 min at 12,000 × g), the supernatant was collected and the soil-bead mixture was extracted a second time by
resuspension in 700 µl of sodium phosphate buffer. Protein and debris
were precipitated from the supernatant by adding 0.4 volume of 7.5 M
ammonium acetate and incubating the mixture on ice for 5 min. After
centrifugation at 12,000 × g for 3 min, nucleic acids
were precipitated by adding 0.7 volume of isopropanol and centrifuging
the mixture at 12,000 × g and 4°C for 45 min.
Subsequently, the DNA pellet was washed with 70% ethanol at 4°C and
dried under vacuum. Finally, DNA was resuspended in 200 µl of
Tris-EDTA buffer (10 mM Tris base, 1 mM EDTA [pH 8]).
Removal of humic acids. The forest soil DNA extracts were dark brown and contained large amounts of humic acids. The humic acids were removed with acid-washed polyvinyl-polypyrrolidone (PVPP) (Sigma-Aldrich Chemie GmbH, Steinheim, Germany) in spin columns (Bio-Rad, Munich, Germany) modified from the method of Holben et al. (21). The spin columns were filled with 2 ml of PVPP, which had been equilibrated and suspended in Tris-EDTA (pH 8). The PVPP columns were packed and dried by centrifugation (375 × g for 1 min) just prior to loading. About 150 µl of the brown humic acid-containing soil DNA extract was loaded onto the column and centrifuged. The purified DNA solution was clear and colorless and readily amplifiable by PCR.
The concentration and purity of the DNA solutions were determined by measurement of absorption at 260 and 280 nm, after 1:10 dilution in H2O, using a GeneQuant spectrophotometer (Pharmacia Biotech, Uppsala, Sweden). For PCR amplification, DNA aliquots at a standardized DNA concentration of 1 ng µl1 were used.
PCR amplification. For PCR amplification, we used a universal SSU rRNA-based primer set, targeting all life, and the functional primer set pmoA (19).
PCR buffer (20 mM Tris-HCl [pH 8.3], 50 mM KCl), 1 U and 0.5 U of AmpliTaq DNA polymerase for the pmoA and the universal primer set, respectively (Perkin-Elmer Applied Biosystems, Weiterstadt, Germany), 0.5 µM each primer, and 100 µM each deoxynucleoside triphosphate (Amersham Life Science, Braunschweig, Germany) were added to a total reaction volume of 50 µl at 4°C. For the pmoA amplification, MasterAmp 2× PCR premix F containing 100 mM Tris-HCl (pH 8.3), MgCl2, 400 µM each deoxynucleoside triphosphate, and the PCR enhancer betaine (Epicentre Technologies, Madison, Wis.) were added to the reaction solutions. The same template concentration (1 to 5 ng µl1) was always used in a set
of PCR amplifications from the same soil core. Amplifications were
started by placing cooled (4°C) PCR tubes immediately into the
preheated (94°C) thermal block of a Mastercycler Gradient
thermocycler (Eppendorf, Hamburg, Germany). The thermal cycling
profiles consisted of touchdown programs with an initial denaturation
of 3 min at 94°C followed by 30 cycles of 30 s at 94°C,
30 s at the annealing temperature, and 45 s at 72°C, with 5 min at 72°C for the last cycle. The annealing temperature decreased
from 62 to 55°C and from 60 to 50°C in 0.5°C steps for the pmoA
and universal primer sets, respectively.
Aliquots (5 µl) of PCR products were analyzed by electrophoresis on
3% agarose gels, stained with ethidium bromide, and quantified densitometrically. The gels were destained in water for 30 min. For
calibration, the Smart-Ladder DNA mass and size ruler (Eurogentec, Seraing, Belgium) was used (calibration coefficient of all analyses, r > 0.9). The gels were photographed with an imaging
system (MWG Biotech, Ebersberg, Germany), and DNA bands were analyzed
with RFLP-scan software (CSP Inc., Billerica, Belgium).
DGGE. DGGE was carried out as described previously in detail (19). PCR products were separated using a DCode System (Bio-Rad) on 1-mm-thick polyacrylamide gels (6.5% [wt/vol] acrylamide-bis acrylamide [37.5:1] [Bio-Rad]) prepared with and electrophoresed in 0.5× TAE (pH 7.4) (0.04 M Tris base, 0.02 M sodium acetate, 1 mM EDTA) at 60°C and constant voltage. A denaturing gradient of 35 to 80% and 35 to 70% was used for the pmoA and Universal PCR products, respectively. A denaturing gradient of 80% (vol/vol) denaturant corresponded to 6.5% acrylamide, 5.6 M urea, and 32% deionized formamide. The gels were poured onto GelBond PAG film (FMC Bioproducts, Rockland, Maine) to avoid gel distortion. The gels were stained with 1:50,000 (vol/vol) SYBR-Green I (Biozym, Hessisch-Oldendorf, Germany) for 30 min and scanned with a Storm 860 PhosphorImager (Molecular Dynamics, Sunnyvale, Calif.). The scanned DGGE gels were digitally enhanced with Photoshop 5.0 (Adobe Systems Inc.) to improve graphic resolution of the figures.
For further analysis, the DGGE bands were visualized in the SYBR Green I-stained gels with blue light (
> 400 nm) using a Dark Reader
transilluminator (Clare Chemical Research, Ross on Wye, United
Kingdom). Individual DGGE bands were then excised, reamplified, and
reanalyzed by DGGE to verify band purity, as described recently
(19).
Sequencing of DGGE bands. Reamplified PCR products of excised DGGE bands were purified using the EasyPure DNA purification kit (Biozym, Hessisch-Oldendorf, Germany). The concentration and purity of the PCR products were determined by measuring the absorption at 260 and 280 nm of a 1:20 dilution in H2O with a GeneQuant spectrophotometer (Pharmacia Biotech). Sequencing reactions were performed using the ABI Dye Terminator cycle-sequencing kit (Perkin-Elmer Applied Biosystems) as specified by the manufacturer. Cycle-sequencing products were purified from excess dye terminators and primers using Microspin G-50 columns (Pharmacia Biotech, Freiburg, Germany) and analyzed with an ABI 373 DNA sequencer (Perkin-Elmer Applied Biosystems).
Sequences were analyzed using the Lasergene software package (DNASTAR, Madison, Wis.). The nucleotide and inferred amino acid sequences of the gene fragments of pmoA were manually aligned with sequences retrieved from the GenBank database. SSU rDNA sequences were aligned and phylogenetically placed with the ARB software package (http://www.biol.chemie.tu-muenchen.de/pub/ARB). The partial SSU rDNA sequences were added to a validated and optimized tree of complete 16S rDNA sequences while keeping the overall topology constant (30). On the nucleic acid level, evolutionary distances between pairs of sequences were calculated by using the Jukes-Cantor and Felsenstein equations (15, 25) provided in the ARB package. Phylogenetic trees were constructed by using distance matrix and maximum-parsimony methods supplied by the ARB software package.Nucleotide accession numbers. The sequences of pmoA gene fragments and of the SSU rDNA fragments of excised DGGE bands have been deposited in GenBank under accession numbers AF200726 to AF200729 and AF200730 to AF200734, respectively.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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CH4 consumption rates by forest soil.
In situ
CH4 mixing ratios decreased with soil depth both in winter
and in summer (Fig. 1). In winter, the
CH4 mixing ratios showed a quasi-linear decrease with soil
depth (Fig. 1A). The scattering of the data was too large to determine
the zone of CH4 oxidation with certainty, but it was
probably at the lower end of the profile. In summer, the in situ
profile showed a quasi-linear decrease in CH4 mixing ratios
from 0 to about 8 cm deep (Fig. 1B). Below 16 cm deep, the in situ
CH4 mixing ratios remained constant at approximately 0.25 ppmv CH4. The shape of the vertical CH4 profile
indicated CH4 oxidation at about 8 to 15 cm deep. The
forest soil was always a net sink for atmospheric CH4,
exhibiting a CH4 consumption rate of 1.00 ± 0.26 mg
m2 day1 (mean ± standard deviation
[SD] of three determinations) in winter. Similar values were reported
previously (1, 29, 38).
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DNA extraction and PCR amplification.
The same soil core
sections that were used for measurement of the CH4
oxidation activity shown in Fig. 2 were also used for DNA extraction.
Up to 10 times more total DNA was extracted from the forest soil in
summer than in winter (Fig. 3A). The
total extractable DNA content of the soil decreased with depth, both in
winter and in summer, but the decrease was more pronounced in summer
(Fig. 3A).
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1 [mean ± SD of 8 determinations]) and
summer (29.5 ± 11.5 ng µl1 [mean ± SD of
13 determinations]) at similar concentrations, indicating that DNA
from all soil layers was readily amplifiable and was not subjected to
different biases caused by humic acids or other PCR-inhibiting substances.
In contrast to the universal primer set, PCR with the primer set pmoA
yielded product concentrations depending on the soil depth and season.
In winter, pmoA was detected only in a well-defined layer
between 6 and 16 cm deep, with the highest PCR product yield being
found between 14 and 16 cm deep (Fig. 3B). No pmoA was
detected in the top 6 cm of soil. In summer, on the other hand,
pmoA was detected and amplified between 0 and 28 cm deep,
with the highest PCR product yield being found between 6 and 8 cm deep
(Fig. 3B). The pmoA gene was detected only in soil layers
with CH4 oxidation activity, indicating that the presence
of the pmoA gene, and thus of methanotrophs, coincided with
the measured activity (Fig. 2A and 3B).
SSU rDNA DGGE community pattern.
The DGGE analysis of PCR
products amplified with the universal primer set was conducted only in
winter (in soil from 0 to 16 cm deep) and revealed a complex banding
pattern, reflecting the high microbial diversity expected for forest
soil (Fig. 4A). The number and intensity
of DGGE bands increased below a soil depth of 2 cm. The DGGE banding
pattern changed at different soil depths, indicating changes of the
bacterial community structure with depth. The intensity of the DGGE
bands indicated the presence of about 11 dominant populations within
the layer of highest diversity (6 to 10 cm deep). Sequence analysis of
five major DGGE bands revealed two populations [MR(UNI)4 and
MR(UNI)5] grouping within the phylum of high-GC gram-positive
bacteria, two populations [MR(UNI)1 and MR(UNI)3] closely related to
the phylum Holophaga-Acidobacterium, and one population
[MR(UNI)2] grouping within the proteobacteria (Fig. 4B). The
latter population [MR(UNI)2] was the most closely related to clones
ms14, ms6, and ms10, which were retrieved from a peat bog with primers
specific for methanotrophs (accession numbers AF111789, AF111787, and
AF111788) (I. McDonald and C. Murrell, personal communication).
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pmoA DGGE community pattern.
The DGGE analysis of
PCR products amplified with the pmoA primer set was performed on soil
samples taken in winter (0 to 16 cm deep) and in summer (0 to 28 cm
deep). In winter, DGGE bands of pmoA products were detected
between 6 and 16 cm deep, and the number of DGGE bands increased from
one to six between 6 and 16 cm deep (Fig.
5A). In the soil layer (10 to 14 cm deep)
with the highest CH4 oxidation rates, the DGGE bands MR2,
MR3, MR4, and MR5 were the most intense.
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proteobacteria and was closely related to the ammonium-oxidizing genus
Nitrosospira. DGGE bands MR2 to MR5 formed a distinct group
of methanotrophs distantly related to common type II methanotrophs ( proteobacteria). These sequences were very similar and clustered with
pmoA clone sequences (Rold 1, 3, and 5; Maine 6, 8, and
RA14) recently retrieved from other forest soils
(23).
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Atmospheric CH4 oxidation has been extensively studied in various upland soils. However, little is known about the methanotrophic community involved. By studying the vertical distribution of both the oxidation of atmospheric CH4 and the molecular characteristics of methanotrophic populations in an acidic forest soil, we detected a methanotrophic community distinct from known type I and II methanotrophs and occurring only in the soil layers which were actively oxidizing atmospheric CH4. The pmoA sequences of this methanotrophic community clustered distantly from the known methanotrophs and grouped with pmoA clone sequences recently retrieved from other forest soils involved in atmospheric CH4 consumption (23).
The forest soil examined in this study consumed atmospheric CH4 at all times. The soil showed a quasi-linear decrease of CH4 with depth, and, as reported for other forest soils, maximal CH4 consumption occurred in subsurface soil layers (1, 29, 33, 38). In winter, no CH4 consumption was measured in the surface layer (0 to 6 cm deep) and no methanotrophs were detected by DGGE with pmoA. One reason for the missing CH4 consumption might be the high water content in winter, which exceeded saturation (>100% WHC). Methane was not produced in the soil, in contrast to observations in other upland soils (13, 28). Therefore, atmospheric rather than endogenously produced CH4 was the only substrate for CH4-oxidizing bacteria in this forest soil.
The inhibition of CH4 oxidation in surface soil was often
attributed to high NH4+ concentrations
(16, 26, 36). The primer set pmoA also amplifies the
amoA gene of ammonium oxidizers coding for the subunit
of ammonium monooxygenase (22). In this forest soil
population, MR1, closely related to Nitrosospira, was
detected below the maximal CH4 consumption zone, while no
ammonium oxidizers were detected near the surface. The presence of
ammonium oxidizers below the soil layer of maximal CH4
oxidation activity raised the question whether ammonium oxidizers were
sustained by nitrification. However, the NH4+
concentrations were not inhibitory to the methanotrophs. Recently, NH4+ was actually identified as a prerequisite
for CH4 oxidation in rice field soil (8).
The increase of total extractable DNA indicated a stimulation and proliferation of all life from January (winter) to July (summer). However, while the total DNA concentration decreased with depth, the pmoA PCR product concentration and the number of methanotrophic populations (i.e., the number of DGGE bands) increased. The methanotrophic community expanded its presence from a well-defined subsoil layer in winter to the entire soil core in summer. It should be noted, however, that the presence of these populations does not necessarily mean that they were active. The MR2 band was present in all depth layers, indicating that MR2 was the most widely distributed methanotrophic population. In the soil layers with highest CH4 oxidation activity, populations MR2, MR3, and MR5 were prevalent over MR4, whereas MR4 and the putative ammonium oxidizer MR1 appeared in the soil layers below those with the maximal oxidation activity. This indicated that the methanotrophic populations were probably adapted to the conditions at a particular depth, each occupying its own ecological niche. The DGGE community pattern itself and the sequences detected were the same in winter and summer.
The most frequently detected bacteria of the forest soil bacterial
community belong to the phyla Holophaga-Acidobacterium, high-GC gram-positive bacteria, and proteobacteria. The proteobacteria DGGE band appeared only in soil layers with
CH4 consumption and was most closely related to clone
sequences (ms10, ms8, and ms14) retrieved with primers specific for
methanotrophs from a peat bog (McDonald and Murrell, personal
communication). However, there is no direct evidence that any of the
populations detected by 16S rDNA-based DGGE indeed belonged to the
methanotrophs. These SSU rDNA sequences, as well as the pmoA
sequences, were distantly related to type II methanotrophs. Recently,
phospholipid fatty acid (PLFA) analysis and sequence comparison of
pmoA in various soils also indicated novel methanotrophs
distantly related to type II methanotrophs and proteobacteria
(23). Although, the correspondence between SSU rDNA and
pmoA data might be coincidental, it allows the hypothesis
that the proteobacteria population detected with the universal
primer set and the methanotrophic populations detected with the pmoA
primer set are the same. The proteobacteria population belonged to
the most abundant bacteria in soil layers with CH4
oxidation. The novel sequences obtained by pmoA PCR and DGGE were found
in the same soil layers. Finally, cloning results in other forest soils
indicated that the same novel pmoA sequences were
ubiquitously distributed in forest soils (23).
Process data and kinetic properties of CH4 consumption have
repeatedly suggested that novel methanotrophs are responsible for
atmospheric CH4 consumption (5, 6, 33). The
novel pmoA sequences strongly support the existence of yet
unknown methanotrophs involved in atmospheric CH4
consumption, as postulated by Bender and Conrad (6).
Radioactive labeling studies showed that high-affinity methanotrophs in
forest soils contain unusual i17:0, a17:0 and 17:1
8c PLFAs whereas
known methanotrophs contain 18:18c and 16:18c PLFAs
(34).
Enrichments with high CH4 concentrations have resulted in the isolation of common methanotrophs from neutral environments. Molecular data from neutral soils and at high CH4 concentrations demonstrate the presence of common type I and II methanotrophs (19, 24, 32). A novel methanotrophic strain, S6, closely related to the genus Beijerinckia, was isolated from an acidic blanket peat, also by using high CH4 concentrations (12). It is unknown whether this strain, which contains only a soluble methane monooxygenase (sMMO), is able to oxidize CH4 with high affinity (nanomolar Km). However, enrichments under low CH4 concentrations resulted in the isolation of the high-affinity type II methanotrophic strain LR1 from neutral soil (14).
In summary, all these observations suggest that known type I and II methanotrophs prevail in neutral environments, preferentially at high CH4 concentrations, but are able to adapt to high-affinity kinetics (type II methanotrophs). However, unknown high-affinity methanotrophs seem to prevail in acidic environments where atmospheric CH4 is oxidized. The missing link in the information is the isolation of a bacterium that combines all the molecular evidence, including unusual pmoA sequence and PLFA pattern, with high-affinity CH4 oxidation kinetics. Forest soils tend to be acidic, and the unknown, high-affinity methanotrophs seem to thrive at a low pH and might therefore have escaped isolation in neutral media (5, 11). This new knowledge about the wide distribution of novel acidicophilic methanotrophs in forest soil, together with the molecular tools at hand, should facilitate the enrichment and isolation of the bacterium which constitutes the missing link.
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ACKNOWLEDGMENTS |
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We thank Bianca Wagner, Sonja Fleissner, and Axel Fey for excellent technical assistance.
This work was supported by grant BIO-4-CT-960419 from the European Commission.
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FOOTNOTES |
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* Corresponding author. Mailing address: Max-Planck-Institut für terrestrische Mikrobiologie, Karl-von-Frisch-Strasse, 35043 Marburg, Germany. Phone: 49-6421-178801. Fax: 49-6421-178809. E-mail: conrad{at}mailer.uni-marburg.de.
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REFERENCES |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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