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Applied and Environmental Microbiology, June 2006, p. 3901-3907, Vol. 72, No. 6
0099-2240/06/$08.00+0     doi:10.1128/AEM.02779-05
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Estimating High-Affinity Methanotrophic Bacterial Biomass, Growth, and Turnover in Soil by Phospholipid Fatty Acid ¹³C Labeling

P. J. Maxfield,¹ E. R. C. Hornibrook,² and R. P. Evershed^1*

Organic Geochemistry Unit, Bristol Biogeochemistry Research Centre, School of Chemistry, University of Bristol, Cantock's Close, Bristol BS8 1TS, United Kingdom,¹ Department of Earth Sciences, Bristol Biogeochemistry Research Centre, University of Bristol, Wills Memorial Building, Queen's Road, Bristol BS8 1RJ, United Kingdom²

Received 24 November 2005/ Accepted 20 March 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
A time series phospholipid fatty acid (PLFA) ¹³C-labeling study was undertaken to determine methanotrophic taxon, calculate methanotrophic biomass, and assess carbon recycling in an upland brown earth soil from Bronydd Mawr (Wales, United Kingdom). Laboratory incubations of soils were performed at ambient CH₄ concentrations using synthetic air containing 2 parts per million of volume of ¹³CH₄. Flowthrough chambers maintained a stable CH₄ concentration throughout the 11-week incubation. Soils were analyzed at weekly intervals by gas chromatography (GC), GC-mass spectrometry, and GC-combustion-isotope ratio mass spectrometry to identify and quantify individual PLFAs and trace the incorporation of ¹³C label into the microbial biomass. Incorporation of the ¹³C label was seen throughout the experiment, with the rate of incorporation decreasing after 9 weeks. The ¹³C values of individual PLFAs showed that ¹³C label was incorporated into different components to various extents and at various rates, reflecting the diversity of PLFA sources. Quantitative assessments of ¹³C-labeled PLFAs showed that the methanotrophic population was of constant structure throughout the experiment. The dominant ¹³C-labeled PLFA was 18:17c, with 16:15 present at lower abundance, suggesting the presence of novel type II methanotrophs. The biomass of methane-oxidizing bacteria at optimum labeling was estimated to be about 7.2 x 10⁶ cells g^–1 of soil (dry weight). While recycling of ¹³C label from the methanotrophic biomass must occur, it is a slower process than initial ¹³CH₄ incorporation, with only about 5 to 10% of ¹³C-labeled PLFAs reflecting this process. Thus, ¹³C-labeled PLFA distributions determined at any time point during ¹³CH₄ incubation can be used for chemotaxonomic assessments, although extended incubations are required to achieve optimum ¹³C labeling for methanotrophic biomass determinations.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Increases in anthropogenic CH₄ emissions have resulted in an increase in the atmospheric methane concentration from 0.75 to 1.8 parts per million of volume (ppmv) in the past 200 years (5, 33). The major terrestrial CH₄ sink is the aerobic oxidation of methane by soil microorganisms, which accounts for approximately 5% of the total sink or 30 Tg year^–1 (21). However, there is a high degree of uncertainty in this value, with estimates ranging from 15 to 45 Tg year^–1 (39).

The activity of soil methanotrophic biomass has primarily been assessed indirectly due to the unculturable nature of high-affinity methanotrophs (11, 16). Determinations of the methane oxidation potential of different soils have shown how this activity varies with changing environmental influences, such as temperature and moisture (3, 38), soil type (6, 35), and land use (20, 27).

New techniques have recently emerged to study bacteria in situ without the need for laboratory cultures to be established. These include gene probe-based methods, such as fluorescent in situ hybridization and PCR targeting pmoA, mmoX, mxaF, and nifH (25), with a significant development being stable isotope probing (SIP) (19, 24, 26, 29). In SIP, DNA and RNA are used as specific bacterial indicators, but quantitative assessments of specific microbial groups are not straightforward in complex environments such as soils.

An alternative molecular approach to the study of high-affinity methanotrophs is to incubate soils with isotopically labeled methane and then assess the activities of methanotrophs through the determination of membrane lipids. Studies have demonstrated uptake of ¹⁴C-radiolabeled methane into different fractions of bacterial biomass and phospholipid fatty acids (PLFAs) (17, 30, 31), and ¹³C stable isotope-tracer studies have been conducted using gas chromatography/combustion/isotope ratio mass spectrometry (GC-C-IRMS), which allows precise ¹³C values to be determined on individual membrane lipid components at high sensitivities (7, 10). The majority of ¹³C-PLFA labeling studies of methanotrophic bacteria have involved incubating soil with ¹³CH₄ in sealed containers (static flux), with the addition of more ¹³CH₄ at intervals to restore the initial headspace concentration. At the end of the incubation period, soils are analyzed for ¹³C-PLFAs (or other membrane lipids) of methanotrophic origin using GC-C-IRMS (7, 10, 12, 22). A shortcoming of using static flux chambers is the difficulty of maintaining constant methane concentrations throughout the experiment; high variability of the CH₄ concentration is undesirable for quantitative studies since this may lead to instability in the methanotrophic community.

In this study a flowthrough incubation chamber was used that allowed long-term atmospheric concentration (2 ppmv) ¹³CH₄ incubations of soils in order to derive a stable, fully ¹³C-labeled methanotrophic population. This enabled the biomass of the active methanotrophic community to be determined via quantification of the ¹³C-labeled PLFAs over a time course.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Field site.
Soils used in this investigation were sampled from the Institute of Grassland and Environmental Research upland research station at Bronydd Mawr (Wales, United Kingdom). The site was part of a low-input farming study and had received no human interference for 13 years. The grassland plot was unfertilized and ungrazed and displayed extensive vegetation growth. Because the site is an upland grassland, the soil profile was relatively shallow and had been previously characterized by the soil survey of Great Britain (32). The soil is classified as a well-drained brown earth, and with bedrock 80 cm from the ground surface, it is a fine loamy soil falling under the international soil taxonomy class of an Entisol.

Soil sampling and incubation.
Soil cores were collected to a depth of 35 cm from duplicate plots. All soil samples were stored in jars at 4°C until analysis. Prior to analysis soils were sieved (2 mm) to achieve homogeneity and remove any large stones. Soils for methane oxidation rate analysis were weighed (10 g) into serum bottles (120 ml), sealed with butyl rubber septa, and stored at 20°C for 24 h in an incubator. Headspace CH₄ concentrations were then monitored over a 5-h period by GC, and methane oxidation rates were calculated by linear regression analysis. All serum bottles were slightly overpressurized to prevent external gas from leaking into the bottle, and any pressure changes were factored into CH₄ uptake calculations.

Soil from a depth of 5 cm was selected for the ¹³CH₄ incubation due to its high methane oxidation rate (0.04 nM CH₄ g^–1 dry weight [dwt] of soil, h^–1). Soil samples for ¹³CH₄ incubation were weighed (10 g) into 5-cm-diameter petri dishes and placed into an incubation chamber (63 liters; Perspex) sealed at the base via immersion in a water-filled tray to prevent gas leakage. The water in the base of the chamber maintained atmospheric moisture saturation and prevented samples from drying. Double-distilled water was added periodically (at ca. 2-week intervals) to the soils during incubation to maintain moisture levels. ¹³CH₄ (2 ppmv) premixed in synthetic air was passed through the chamber at a rate of 44 ml min^–1 in order to renew the headspace every 24 h. Each soil sample was incubated in triplicate with incubation times ranging from 2 to 11 weeks. A second, identical set of "control" soils was incubated in a separate building under ambient CH₄ (2 ppmv) levels. All incubations were carried out in temperature-controlled rooms at 20°C. Throughout the experiment selected soil samples were removed for isotope analysis and monitored by GC headspace analysis to verify that methane oxidation activity was being maintained.

PLFA analysis.
Following incubation, all soil samples were freeze-dried, homogenized by grinding, and extracted using a modified Bligh-Dyer monophase solvent system (4). Briefly, soil (2 g dwt) was extracted using Bligh-Dyer solvent containing water buffer (0.05 M KH₂PO₄; pH 7.2)-chloroform-methanol in a ratio of 4:5:10 (vol/vol/vol). After the addition of solvent (3 ml), the mixture was sonicated (15 min) and centrifuged (at 3,200 rpm for 5 min). The supernatant was decanted, and the residue was extracted as above (three times). The extracts were combined and evaporated under nitrogen to yield the total lipid extract. The total lipid extract was separated into three fractions based on polarity using a 3-ml STRATA aminopropyl-bonded silica cartridge (Phenomenex). Fractions were eluted sequentially with 2:1 (vol/vol) dichloromethane-isopropanol (8 ml; yielding neutrals), 2% acetic acid in diethyl ether (12 ml; free fatty acids), and methanol (8 ml; polar fraction including phospholipids). The PLFA components of the polar fraction were released by saponification (1 ml of 0.5 M methanolic NaOH for 1 h at 80°C), and an internal standard was added (10 µl of 0.1 mg ml^–1 solution of C₁₉ n-alkane in hexane). The PLFAs were then methylated using a 14% (vol/vol) BF₃-methanol solution (100 µl; Aldrich, United Kingdom) (1 h at 80°C) and extracted with chloroform (three samples of 2 ml each). The phospholipid ester-linked fatty acid methyl esters (PLFAMEs) were dissolved in hexane for analysis by GC, GC-MS, and GC-C-IRMS.

Dimethyl disulfide derivatives of monounsaturated fatty acids were prepared to facilitate determination of double-bond positions and geometries in monounsaturated fatty acids. A 100-µl aliquot of the methyl esters in dichloromethane was added to 0.25 M I₂/diethyl ether (100 µl) and dimethyl disulfide (1 ml) and then heated (at 60°C for 24 h in the dark). Excess I₂ was removed by addition of aqueous Na₂S₂O₃ (5%; 2 ml), and the dimethyl disulfide derivatives were extracted with hexane (two times; 2 ml).

Instrumental analysis. (i) GC.
Gas samples from the methane oxidation rate experiments were analyzed isothermally using a Carlo Erba 5300 series GC containing a packed column (Porapak Q; 3 mm by 3 m) at 35°C. Sample introduction was via a 50-µl sample loop, and detection was performed with a flame ionization detector operating with ultrapure air (BOC specialty gases) to minimize baseline fluctuations.

PLFAs were analyzed on a Carlo Erba 5300 series GC. Sample introduction was via an on-column injector, and the detector was a flame ionization detector. FAME separation was achieved using a Varian Factor Four VF23MS column (60 m by 0.32 µm [internal diameter]; 0.15-µm film thickness). The carrier gas was hydrogen, and the oven temperature was programmed from 50°C (held for 2 min) to 100°C at 15°C min^–1, from 100 to 220°C at 4°C min^–1, from 220 to 240°C (held for 5 min) at 15°C min^–1.

(ii) GC-MS.
GC-MS analyses were conducted using a Thermofinnigan Trace GC-MS system. The column and temperature program details were the same as those described in the above section. The ion source and the transfer line were maintained at 200 and 260°C, respectively.

(iii) GC-C-IRMS.
GC-C-IRMS analyses of PLFAMEs were carried out using a Thermofinnigan Delta Plus_XL IRMS (electron ionization, 100-eV electron voltage and 1-mA electron energy; three faraday cup collectors for m/z 44, 45, and 46) instrument linked to a Hewlett Packard 6890N GC via a Thermofinnigan version 3 combustion interface with a copper oxide and platinum catalyst maintained at 850°C. Water removal was via a Nafion membrane, and the column and temperature program details were the same as those detailed in the above section but with helium as the carrier gas. Reference CO₂ of known isotopic composition was used for sample calibration and introduced directly into the source three times at the start and end of each run. Each sample was run in duplicate to ensure reliable mean ¹³C values.

Isotopic mass balancing.
All ¹³C values were corrected for derivatization using a mass balance equation (equation 1). The ¹³C value of the BF₃-methanol was –41.1 ± 0.1, determined by bulk IRMS.

(1)

where n is the number of carbon atoms, c is the compound of interest, d is the derivatizing agent, and cd is the derivatized compound of interest. The fractional abundance (F) of ¹³C in the control (F_c) and enriched (F_e) PLFAs was used to calculate the concentration of ¹³C incorporated into PLFAs from the total PLFA concentration (equation 2).

(2)

The fractional abundance expresses the amount of ¹³C as a proportion of the total amount of carbon in the PLFA (equation 3).

(3)

Statistical analysis.
Bacterial populations were compared by cluster analysis using SYSTAT, version 7.0. The distribution of PLFAs was used as a measure of similarity or distance. The analysis was performed on Euclidean distances between standardized data using averages.

Top Abstract Introduction Materials and Methods Results Discussion References

 
PLFA profiles.
Table 1 shows the concentrations of the major PLFAs extracted from the Bronydd Mawr soil following incubation of 2 to 11 weeks. Microbial PLFAs are present ranging predominantly from C₁₆ to C₁₈ with the most abundant being 16:0, 18:17c, and 18:19c, but without incubation with ¹³CH₄ and compound-specific stable carbon isotope analysis, it is not possible to determine which PLFAs derive from methanotrophs. During the course of the incubation experiment, the concentrations of the PLFAs increased. After 2 weeks of incubation the most abundant PLFA, 18:17c, displayed a concentration of 51.1 nM g^–1 (dwt) of soil, increasing to 182.5 nM g^–1 (dwt) of soil after 11 weeks.

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TABLE 1. Concentration of major PLFAs extracted from Bronydd Mawr soil at a depth of 5 cm

¹³CH₄ labeling technique.
The clearest way to display changes in compound-specific carbon isotope values is as ¹³C values (¹³C_labeled – ¹³C_control) (Fig. 1). It can be seen that there is a marked and steady incorporation of ¹³C label into selected PLFAs with time: 16:15 and 18:7c show the largest increase in ¹³C values, rising by 50.0 and 38.4, respectively, after 11 weeks; i17:0 and 18:15c displayed the next largest increases in ¹³C values of 15.1 and 9.8, respectively. Standard deviations are relatively low (16:15 excluded) for such a complex biological system. ¹³C label incorporation was slow compared to other studies that have used higher concentrations of CH₄ (12), reaching a maximum ¹³C value of 50 after 11 weeks.

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FIG. 1. 13C values of selected PLFAs incorporating 13C label following incubation with 2 ppmv of 13CH4. The division into graphs A and B enables the PLFAs incorporating lower proportions of 13C label to be seen more clearly. Error bars represent ±1 standard deviation.

Figure 2 compares the concentrations of PLFAs incorporating the ¹³C label throughout the study and indicates that ¹³C-labeled PLFA concentrations increase with time. Converting ¹³C values to absolute concentrations of ¹³C-PLFA (7) provides a true representation of the methanotrophic community PLFA distribution, indicating 18:17c to be the dominant PLFA. Figure 2 also shows the relative abundance of ¹³C-labeled PLFAs for each time point. Within experimental error, the composition of the methanotrophic community, as judged by the constancy of the ¹³C-PLFA distribution, was maintained throughout the experiment.

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FIG. 2. Total concentration of 13C label incorporated into each PLFA in ng of 13C g–1 (dwt) of soil (black bars) following incubation with 2 ppmv of 13CH4 and relative mole percentage abundance (white bars) for the main 13C-labeled PLFAs from 2 to 11 weeks following incubation with 2 ppmv of 13CH4. PLFA analyses were performed on soil samples taken at weekly intervals, and only PLFAs with a relative abundance of >3% are presented. Error bars represent ±1 standard deviation.

Taxonomic identity of high-affinity methane-oxidizing bacteria.
The ¹³C-labeling study shows that several PLFAs are directly derived from high-affinity methane-oxidizing bacteria. A comparison of the ¹³C-labeled PLFA distribution extracted from the Bronydd Mawr soil with published PLFA compositions of pure cultures of methanotrophic bacteria (9, 14) is shown in Fig. 3. The cluster analysis indicates that the bacteria mediating CH₄ oxidation are most closely related to Methylocella palustris (8). The PLFA profiles of the methanotrophs from the Bronydd Mawr soil and M. palustris are both dominated by 18:17c.

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FIG. 3. Comparison of the 13C-labeled PLFA distribution extracted from the Bronydd Mawr soil following incubation with 2 ppmv of 13CH4 with published PLFA compositions of pure cultures of methanotrophic bacteria (9, 14). PLFA compositions used were the mole percentages of the PLFAs of pure cultures and the labeled PLFAs extracted from the soil. A hierarchical tree was produced by cluster analysis performed with the software package SYSTAT, version 7, on Euclidean distances between the standardized data using averages.

Quantification of high-affinity methane-oxidizing bacteria.
By determining the total amount of ¹³C incorporated into the bacterial PLFAs, it is possible to estimate the methanotrophic biomass. Frostegård and Bååth (15) calculated an average value of 1.4 x 10^–17 mol of bacterial PLFA cell^–1. Using this value an estimate of methanotrophic biomass of 5.6 x 10⁶ to 8.8 x 10⁶ methanotrophic cell g^–1 (dwt) of soil was obtained. The range of this estimate takes into account experimental errors. Moreover, the upper and lower limits are also dependent on the concentration of ¹³C-labeled PLFAs resulting from recycling of the ¹³C label into the total bacterial biomass. The upper limit of the estimate of 8.8 x 10⁶ methanotrophic cells g^–1(dwt) of soil is calculated from all PLFAs that incorporated ¹³C label (including those not known to be produced by culturable methanotrophs) and, hence, may be a slight overestimate as a result of ¹³C label recycling. In contrast, the estimate of 5.6 x 10⁶ methanotrophic cells g^–1 (dwt) of soil is based on only the major ¹³C-labeled methanotrophic PLFA, namely 18:17c.

Figure 4 plots estimated ¹³C-labeled methanotroph cell numbers at each time point throughout the incubation. Incorporation of ¹³C label was slow initially but increased rapidly between 6 and 9 weeks, before slowing during weeks 9 and 10. The curves derived from both ¹³C-labeled 18:17c and total bacterial ¹³C-labeled PLFAs appeared to plateau at 10 to 11 weeks. This indicates that the microbial population was at or close to a steady state, with the majority of methanotrophic bacteria having incorporated the ¹³C label.

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FIG. 4. Increase in number of 13C-labeled methanotrophic bacterial cells throughout the 13CH4 incubation. The plot of 18:17c (open squares) represents the main labeled PLFA and is unlikely to include recycled 13C, whereas the plot of all 13C-labeled PLFAs (filled squares) may include up to ca. 10% recycled 13C label. Error bars represent ±1 standard deviation. Trend lines indicate the moving average of two data points.

Top Abstract Introduction Materials and Methods Results Discussion References

 
¹³CH₄ labeling technique.
The flowthrough incubation system employed in this investigation is clearly an advantage over the static flux chambers used in previous ¹³C-labeling studies of high-affinity methanotrophs (7, 10, 12, 22). The degree of ¹³C label incorporation was relatively low, which reduces problems with ¹³C "carryover effects" into closely eluting PLFAs. Mottram and Evershed (28) observed that ¹³C-enriched FAMEs possessing ¹³C values of 500 created a significant within-run carryover effect into unlabeled FAMEs eluting immediately after ¹³C-enriched components. While not a significant problem in qualitative ¹³C-labeling studies, this effect may introduce significant errors in quantitative studies involving ¹³C labeling.

A second advantage of low ¹³C labeling is the ability to assess incorporation at a ¹³CH₄ concentration close to the ambient atmospheric methane concentration. This is considerably more representative of a natural soil system than the 10- to 50-ppmv concentrations typically used in static chamber incubations (7, 12, 22) and the elevated concentrations utilized in DNA-based SIP studies (19, 24, 29). Radajewski et al. (29) estimated that gravimetrically separated ¹³C-DNA fractions contained 65 to 100% ¹³C. To achieve this magnitude of ¹³C label incorporation, artificially high ¹³CH₄ concentrations are applied, with most studies utilizing ¹³CH₄ concentrations above 1% (vol/vol). RNA-based SIP is more sensitive than DNA-based SIP and could potentially characterize active high-affinity methanotrophs following incubations at ambient ¹³CH₄ concentrations, if limitations associated with extracting RNA from environmental samples can be overcome (25).

In this experiment the maximum incubation time was 11 weeks, which is a substantially longer incubation period than has been used in other investigations (7, 10, 12, 22); the results obtained indicate that even after this period of time the microbial population remains fully active in oxidizing methane. After 9 weeks of incubation, it would appear that the high-affinity methanotroph community has reached a steady state in terms of growth and ¹³C turnover, suggesting that methanotrophic population estimates made at that point are representative of the active methanotrophic bacteria in the system.

Although the incubation period was longer than that used in previous studies (7, 10, 12, 22), it appeared that recycling of ¹³C label among other microorganisms was negligible. One of the main aims of this study was to monitor the incorporation of ¹³CH₄ over time to investigate whether any significant turnover of the ¹³C label into nonmethanotrophic PLFAs occurred. If PLFAs initially show no ¹³C label incorporation but then incorporate ¹³C label after an extended period of incubation, it is likely that the microbes from which they derive are not obtaining carbon for biosynthesis directly from CH₄ but through utilization of other substrates containing ¹³C label. Figure 2 indicates that, within experimental error, the ¹³C-labeled PLFA distribution remains consistent throughout the incubation period, thereby excluding the possibility of a significant proportion of ¹³C-labeled PLFAs resulting from recycling of the ¹³C label within the wider microbial biomass. Considerable care was taken when designing the ¹³CH₄ flowthrough incubation system to introduce low concentrations of ¹³CH₄ to the system and extract all respired gases (including ¹³CO₂), thereby limiting any ¹³C-labeled PLFAs derived through ¹³CO₂ utilization by chemoautotrophic bacterial communities. Moreover, controlling the concentrations of ¹³CH₄ at 2 ppmv, i.e., close to the atmospheric concentration, ensured that only high-affinity methanotrophs would be active, thus minimizing the production of unnaturally high concentrations of ¹³C-labeled metabolites or bacterial necromass.

Quantification of methanotrophic bacterial biomass.
By employing the PLFA concentration per bacterial cell derived by Frostegård and Bååth (15), we estimated the population of methanotrophic bacteria at ca. 7.2 x 10⁶ cells g^–1 (dwt) of soil (range, 5.6 x 10⁶ to 8.8 x 10⁶ cells g^–1). Incorporation of the ¹³C label had slowed toward the end of the 11-week incubation, indicating that the microbial biomass had reached, or was close to, a steady state. When considering this methanotrophic biomass estimate, it is important to note that the conversion factor used in the calculation is based on a mean PLFA concentration for a wide range of bacteria (15). We are currently improving this estimate by deriving a specific PLFA concentration for methanotrophic bacteria.

Few studies have made an assessment of methanotrophic biomass in terrestrial environments. Bender and Conrad (1, 2) used a direct most probable number technique to count methanotrophic bacteria from a wide range of different environments. The technique works well at high CH₄ concentrations but is unsuitable for enumerating high-affinity methanotrophs that oxidize CH₄ at ambient concentrations. Horz et al. (18) also used a most probable number technique to quantify the methane-oxidizing bacteria in a German wet meadow soil and estimated the most abundant methanotroph at 10⁵ to 10⁷ cells g^–1 (dwt) of soil. Kolb et al. (23) utilized quantitative PCR to calculate the biomass of methanotrophic bacteria in acidic and neutral forest soils. They detected both culturable and nonculturable methanotrophs in forest soils in the range of 10⁶ gene copies g^–1 (dwt) of soil. The number of cells is lower than this as at least two pmoA gene copies can be expected per cell (34), and unrepresentative discrepancies can be exaggerated by bias introduced during PCR amplification (37). Sundh et al. (36) estimated the cell numbers of methanotrophic bacteria in boreal peatlands by focusing on two specific PLFAs believed to be unique to methanotrophic bacteria, namely 16:18 and 18:18; the total number of methanotrophic bacteria estimated ranged from 0.3 x 10⁶ to 51 x 10⁶ cells g^–1 of wet peat (36), although it is unknown whether methanotrophic bacteria are the only bacteria that exclusively produce 8 monounsaturated PLFAs.

Our estimate of methanotrophic biomass appears low compared to the total bacterial biomass of 5 x 10¹⁰ cells, but it is reasonable when considering the widely differing concentrations of substrates available to the total soil microbial community compared to the concentration of CH₄ available to the methanotrophic bacteria. Roslev et al. (30) calculated theoretical microbial biomass production with CH₄ as the sole carbon source and determined that the potential for population growth solely utilizing atmospheric methane is limited. They estimated a production of ca. 8 x 10⁶ bacteria cm^–3 day^–1. The number of methanotrophic bacteria calculated in the present study is somewhat lower than their estimated daily production value, but CH₄ is a minimal carbon source in aerobic mineral soils, and there are many other more extensively bioavailable carbon substrates.

The ¹³C-labeled PLFA distribution obtained herein for the high-affinity methane-oxidizing bacteria present in the Bronydd Mawr soils shows that they are similar to known type II methanotrophs, agreeing with our previous studies of a range of northern European soils (10, 12). The PLFA distribution obtained in this study, particularly the high abundance of 18:17c, indicates that the Bronydd Mawr methanotrophs are a novel species closely related to M. palustris (Fig. 3) (9). PmoA gene data for the active methanotrophic bacteria in the Bronydd Mawr soil would improve the accuracy of this taxonomic assignment, but this was not the primary aim of this investigation. Overall, indications are that although there are similarities in the high-affinity methanotrophic populations active in soils, there are also some differences between the populations operating in different environments. Other studies have shown that high-affinity methanotrophs are often closely related to known type II low-affinity methanotrophs (10, 12), which could be due to the low-substrate conditions that high-affinity methanotrophs tolerate with very low CH₄ concentrations. Type II methanotrophs have been found in environments that are more nutrient limited than those typically inhabited by type I methanotrophs, such as oxygen-poor layers of landfill-cover soils (13).

Conclusions.
We have demonstrated herein a new flowthrough ¹³C-labeling approach which allows the simultaneous determination of both methanotrophic biomass and taxon using PLFA ¹³C labeling. It has been shown that an Alphaproteobacteria-specific PLFA (18:17c) incorporated ¹³C label over a period of 11 weeks. We have determined the active high-affinity methanotrophic biomass based on concentrations of ¹³C-labeled PLFAs and shown that there is no substantial turnover or incorporation of the ¹³C label into nonmethanotrophic PLFAs.

 
We thank I. D. Bull and R. Berstan for technical assistance. We are grateful for the use of the NERC Life Sciences Mass Spectrometry Facility, and we thank the Institute of Grassland and Environmental Research for field site access at Bronydd Mawr.

The research facility used in this study at the Bristol Biogeochemistry Research Centre was funded in part from a recent major infrastructure grant provided by the United Kingdom Joint Higher Education Funding Council for England and Office of Science and Technology Science Research Investment Fund and by the University of Bristol. This work was partially supported by a Royal Society grant (574006.G503/24026/SM) to E.R.C.H. This investigation was performed while P.J.M. was in receipt of an NERC studentship.

 
* Corresponding author. Mailing address: Bristol Biogeochemistry Research Centre, School of Chemistry, University of Bristol, Cantock's Close, Bristol BS8 1TS, United Kingdom. Phone: 44 117 928 7671. Fax: 44 117 929 3746. E-mail: r.p.evershed{at}bris.ac.uk.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Bender, M., and R. Conrad. 1992. Kinetics of CH₄ oxidation in oxic soils exposed to ambient air or high CH₄ mixing ratios. FEMS Microbiol. Ecol. 101:261-270.[CrossRef]
2
2. Bender, M., and R. Conrad. 1994. Methane oxidation activity in various soils and fresh-water sediments—occurrence, characteristics, vertical profiles, and distribution on grain-size fractions. J. Geophys. Res. 99:16531-16540.[CrossRef]
3
3. Billings, S. A., D. D. Richter, and J. Yarrie. 2000. Sensitivity of soil methane fluxes to reduced precipitation in boreal forest soils. Soil Biol. Biochem. 32:1431-1441.[CrossRef]
4
4. Bligh, E. G., and W. J. Dyer. 1959. A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37:911-917.
5
5. Blunier, T., J. A. Chappellaz, J. Schwander, J. M. Barnola, T. Desperts, B. Stauffer, and D. Raynaud. 1993. Atmospheric methane, record from a Greenland ice core over the last 1000 years. Geophys. Res. Lett. 20:2219-2222.
6
6. Boeckx, P., O. VanCleemput, and I. Villaralvo. 1997. Methane oxidation in soils with different textures and land use. Nutr. Cycle Agroecosyst. 49:91-95.[CrossRef]
7
7. Boschker, H. T. S., S. C. Nold, P. Wellsbury, D. Bos, W. de Graaf, R. Pel, R. J. Parkes, and T. E. Cappenberg. 1998. Direct linking of microbial populations to specific biogeochemical processes by ¹³C-labelling of biomarkers. Nature 392:801-805.[CrossRef]
8
8. Bowman, J. P., J. H. Skerratt, P. D. Nichols, and L. I. Sly. 1991. Phospholipid fatty-acid and lipopolysaccharide fatty-acid signature lipids in methane-utilizing bacteria. FEMS Microbiol. Ecol. 85:15-22.[CrossRef]
9
9. Bowman, J. P., L. I. Sly, P. D. Nichols, and A. C. Hayward. 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.[Abstract/Free Full Text]
10
10. Bull, I. D., N. R. Parekh, G. H. Hall, P. Ineson, and R. P. Evershed. 2000. Detection and classification of atmospheric methane oxidising bacteria in soil. Nature 405:175-178.[CrossRef]
11
11. Conrad, R. 1996. Soil microorganisms as controllers of atmospheric trace gases (H₂, CO, CH₄, OCS, N₂O, and NO). Microbiol. Rev. 60:609-664.[Abstract/Free Full Text]
12
12. Crossman, Z. M., P. Ineson, and R. P. Evershed. 2005. The use of ¹³C-labelling of bacterial lipids in the characterisation of ambient methane-oxidising bacteria in soils. Org. Geochem. 36:769-778.[CrossRef]
13
13. Crossman, Z. M., F. Abraham, and R. P. Evershed. 2004. Stable isotope pulse-chasing and compound specific stable carbon isotope analysis of phospholipid fatty acids to assess methane oxidizing bacterial populations in landfill cover soils. Environ Sci. Technol. 38:1359-1367.[Medline]
14
14. Dedysh, S. N. 2002. Methanotrophic bacteria of acidic sphagnum peat bogs. Microbiology 71:638-650.[CrossRef]
15
15. Frostegård, Å., and E. Bååth. 1996. The use of phospholipid fatty acid analysis to estimate bacterial and fungal biomass in soil. Biol. Fertil. Soils 22:59-65.
16
16. Hanson, R. S., and T. E. Hanson. 1996. Methanotrophic bacteria. Microbiol. Rev. 60:439-471.[Abstract/Free Full Text]
17
17. Holmes, A. J., P. Roslev, I. R. McDonald, N. Iversen, K. Henriksen, and J. C. Murrell. 1999. Characterization of methanotrophic bacterial populations in soils showing atmospheric methane uptake. Appl. Environ. Microbiol. 65:3312-3318.[Abstract/Free Full Text]
18
18. Horz, H. P., A. S. Raghubanshi, E. Heyer, C. Kammann, R. Conrad, and P. F. Dunfield. 2002. Activity and community structure of methane-oxidising bacteria in a wet meadow soil. FEMS Microbiol. Ecol. 41:247-257.[CrossRef]
19
19. Hutchens, E., S. Radajewski, M. G. Dumont, I. R. McDonald, and J. C. Murrell. 2004. Analysis of methanotrophic bacteria in Movile Cave by stable isotope probing. Environ. Microbiol. 6:111-120.[CrossRef][Medline]
20
20. Hutsch, B. W. 1998. Tillage and land use effects on methane oxidation rates and their vertical profiles in soil. Biol. Fertil. Soils 27:284-292.[CrossRef]
21
21. Khalil, M. 2000. Sources of methane: an overview, p. 98-111. In M. Khalil (ed.), Atmospheric methane: its role in the global environment. Springer-Verlag, New York, N.Y.
22
22. Knief, C., A. Lipski, and P. F. Dunfield. 2003. Diversity and activity of methanotrophic bacteria in different upland soils. Appl. Environ. Microbiol. 69:6703-6714.[Abstract/Free Full Text]
23
23. Kolb, S., C. Knief, P. F. Dunfield, and R. Conrad. 2005. Abundance and activity of uncultured methanotrophic bacteria involved in the consumption of atmospheric methane in two forest soils. Environ. Microbiol. 7:1150-1161.[CrossRef][Medline]
24
24. Lin, J. L., S. Radajewski, B. T. Eshinimaev, Y. A. Trotsenko, I. R. McDonald, and J. C. Murrell. 2000. Molecular diversity of methanotrophs in Transbaikal soda lake sediments and identification of potentially active populations by stable isotope probing. Environ. Microbiol. 6:1049-1060.
25
25. McDonald, I. R., S. Radajewski, and J. C. Murrell. 2005. Stable isotope probing of nucleic acids in methanotrophs and methylotrophs: a review. Org. Geochem. 36:779-787.[CrossRef]
26
26. Morris, S. A., S. Radajewski, T. W. Willison, and J. C. Murrell. 2002. Identification of the functionally active methanotroph population in a peat soil microcosm by stable-isotope probing. Appl. Environ. Microbiol. 68:1446-1453.[Abstract/Free Full Text]
27
27. Mosier, A. R., J. A. Delgado, V. L. Cochran, D. W. Valentine, and W. J. Parton. 1997. Impact of agriculture on soil consumption of atmospheric CH₄ and a comparison of CH₄ and N₂O flux in subarctic, temperate and tropical grasslands. Nutr. Cycle Agroecosyst. 49:71-83.[CrossRef]
28
28. Mottram, H. R., and R. P. Evershed. 2003. Practical considerations in the gas chromatography/combustion/isotope ratio monitoring mass spectrometry of ¹³C-enriched compounds: detection limits and carryover effects. Rapid Commun. Mass Spectrom. 17:2669-2674.[CrossRef][Medline]
29
29. Radajewski, S., G. Webster, D. S. Reay, S. A. Morris, P. Ineson, D. B. Nedwell, J. I. Prosser, and J. C. Murrell. 2002. Identification of active methylotroph populations in an acidic forest soil by stable isotope probing. Microbiology 148:2331-2342.[Abstract/Free Full Text]
30
30. Roslev, P., N. Iversen, and K. Henriksen. 1997. Oxidation and assimilation of atmospheric methane by soil methane oxidizers. Appl. Environ. Microbiol. 63:874-880.[Abstract]
31
31. Roslev, P., and N. Iversen. 1999. Radioactive fingerprinting of microorganisms that oxidize atmospheric methane in different soils. Appl. Environ. Microbiol. 65:4064-4070.[Abstract/Free Full Text]
32
32. Rudeforth, C. C., R. Hartnup, J. W. Lea, T. R. E. Thompson, and P. S. Wright. 1984. Soils and their use in Wales. Bull. 11. Soil Survey of England and Wales, Harpenden, United Kingdom.
33
33. Stauffer, B., G. Fischer, A. Neftel, and H. Oeschger. 1985. Increase of atmospheric methane recorded in Antarctic ice core. Science 229:1386-1388.[Abstract/Free Full Text]
34
34. Stolyar, S., A. M. Costello, T. L. Peeples, and M. E. Lidstrom. 1999. Role of multiple gene copies in particulate methane monooxygenase activity in the methane-oxidizing bacterium Methylococcus capsulatus Bath. Microbiology 145:1235-1244.[Abstract/Free Full Text]
35
35. Striegl, R. G., T. A. McConnaughey, D. C. Thorstenson, E. P. Weeks, and J. C. Woodward. 1992. Consumption of atmospheric methane by desert soils. Nature 357:145-147.[CrossRef]
36
36. Sundh, I., P. Borga, M. Nilsson, and B. H. Svensson. 1995. Estimation of cell numbers of methanotrophic bacteria in boreal peatlands based on analysis of specific phospholipid fatty-acids. FEMS Microbiol. Ecol. 18:103-112.
37
37. Von Wintzingerode, F., U. B. Gobel, and E. Stackebrandt. 1997. Determination of microbial diversity in environmental samples: pitfalls of PCR-based rRNA analysis. FEMS Microbiol. Rev. 21:213-229.[CrossRef][Medline]
38
38. Whalen, S. C., and W. S. Reeburgh. 1996. Moisture and temperature sensitivity of CH₄ oxidation in boreal soils. Soil Biol. Biochem. 28:1271-1281.[CrossRef]
39
39. Wuebbles, D. J., and K. Hayhoe. 2002. Atmospheric methane and global change. Earth Sci. Rev. 57:177-210.[CrossRef]

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Applied and Environmental Microbiology, June 2006, p. 3901-3907, Vol. 72, No. 6
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