ELSEVIER
FEMS Microbiology
Ecology
18 (1995) 103-l 12
Estimation of cell numbers of methanotrophic bacteria in boreal
peatlands based on analysis of specific phospholipid fatty acids
Ingvar Sundh a-*, Peter Borg; b, Mats Nilsson ‘, Bo H. Svensson a
a Department
of Microbiology.
’ Department
of Chemistry,
Department
Swedish Unirersi~
Swedish Uniwrsity
of Forest Ecology.
Stredish
Received 9 February
ofAgricultural
of Agricultural
Unirersity
Sciences, Box 7025. S-750 07 Uppsala.
Sciences, Bor 7015, S-750 07 Uppsala.
of Agricultural
Sweden
Sweden
Sciences. S-901 83 C/me& Sweden
1995; revised 3 June 1995: accepted 3 July 1995
Abstract
Concentrations
of two phospholipid
were used to estimate
fatty
acids
(PLFAs)
Pearland:
and cell numbers
Methane oxidation;
Methanotrophic
specific
for methane-oxidizing
bacteria
(16:
I ~8 and 18:I ~81,
of this group of bacteria in two Sphagnum-dominated
boreal peatlands.
Concentration ranges of 16: I w8 and 18: I w8 were 0.0-73 and I .O-486 pmol g - ’ of wet peat, respectively. Concentrations
in the peat of each fatty acid were positively correlated with the potential methane oxidation activity (V,,,),
which was used
as an independent estimate of methanotrophic biomass. This correlation
suggests
that the two PLFAs are good biomarkers
for the population of methanotrophic
bacteria in peatlands. Concentrations
of the two PLFAs were transformed to cell
numbers using conversion factors for the cell content of PLFAs, average cell volume and percentage
of cellular dry matter.
The total cell number of methanotrophic bacteria in peat samples from a range of sites and depths ranged between 0.3 and
51 X 10h cells g-’ of wet peat, with similar proportions of type I and type II methanotrophic bacteria in most samples.
Within particular peat profiles, numbers of methanotrophic
bacteria were highest around the level of the water table,
implying that the supplies of methane and oxygen largely determine the biomass distribution of methanotrophic bacteria in
this type of peatlands.
Kewcord.~:
the biomass
bacteria;
Biomass;
1. Introduction
Methane-oxidizing
(methanotrophic)
bacteria are
widespread in nature and play a major role in the
global biogeochemistry
of methane, an important
‘greenhouse’ gas [ 1,2]. They typically occur at the
aerobic/anaerobic
interface in wet environments
such as lake sediments,
rice paddies and water-
* Corresponding
author. Tel.: +46 I8 67 32 10; Fax: +46
67 33 92: E-mail: [email protected].
0168-6496/95/$09.50
SSDI 0168.6496(95)00046-
0
1995 Federation
1
of European
18
Microbiological
Biomarker:
Phospholipid
fatty acid
saturated peat soils [3-61. In these environments,
they utilize methane diffusing from anaerobic zones
and often function as a filter for methane that would
otherwise be emitted to the atmosphere. Many welldrained mineral soils are net consumers of atmospheric methane [7,8], indicating
the presence of
methanotrophic bacteria in these soils as well.
Many different strains of methanotrophic bacteria
have been isolated in pure culture, and based on the
mole percentage of guanine-cytosine
in their DNA,
intracellular membrane arrangement, carbon assimilation pathway, phospholipid fatty acid (PLFA) comSocieties.
All rights reserved
position and some other features, the methanotrophic
bacteria have been classified as type I, type II or type
X [9]. Most of the PLFAs in these bacteria are
monounsaturated
and contain chains of 16 (type I
and Xl or 18 (type II) carbon atoms [lO,l I]. Additionally, in roughly half of the isolates for which the
PLFA composition has been determined, a high proportion of the fatty acids have the double bond in the
very unusual 08 position (i.e. at the 8th carbon atom
from the aliphatic end of the molecule) [ 10,12,13].
This makes these PLFAs potentially useful as specific biomarkers for methanotrophic bacteria in complex environmental
samples. It has been observed
that the concentration
of these unusual fatty acids
increased when a soil [14] and a sandy pond sediment [ 151 were enriched with methane or natural gas.
In this study, the goal was to evaluate the use of
the unusual PLFAs present in methanotrophic bacteria as biomarkers for these bacteria in boreal peatlands. Beside determining the concentrations
in peat
of monounsaturated
phospholipid fatty acids of 16
and 18 carbon length with the double-bond in the w8
position, the potential methane oxidation (i.e. measured with an excess supply of methane) was used as
an independent
estimate of the total biomass of
methanotrophic microorganisms.
2. Materials
and methods
2. I. Description
of sites
Peat samples were collected at a total of eight
sites in two mires in the northern boreal zone in
Sweden. Brief descriptions of the two mires and the
sites of peat sampling are given here. For more
details see Sundh et al. [5,16]. The botanical nomenclature follows Koponen et al. for mosses [17] and
Lid for yascular plants [ 181.
Stor-Amyran (63”44’N, 20”06’E) is an acid mixed
mire. It has an area of 1 km’ and contains both
ombrotrophic
(nutrient supply via deposition only;
relatively nutrient poor) and minerotrophic (nutrients
also supplied via surface0 flow; relatively richer in
nutrients) parts. At Stor-Amyran.
several species of
Sphagnum dominate the bottom layer. Common vascular plants in areas with the water table close to the
vegetation surface include Eriophorum caginatum
and
Drosera
anglica.
whereas
Ox~cocc~s
quadripetalus, Calluna Lulgaris, Andromeda polifolia, Rubus chamaemorus
and Pinus siluestris are
common in drier communities.
Peat samples were
collected at five sites: a wet hollow. a mud-bottom
(both with the water table at the vegetation surface
for most of the growing season), a string, a hummock and one site at the minerotrophic part of the
mire (the last three with water tables at IO-30 cm
depth).
Bjommyran (64”20’N, 18” 18’E) has an area of 2.5
km’. It is a medium-rich tall sedge fen with a large
part consisting of well-developed
string-flark patterns. On the strings, several species of Sphagnum
dominate the bottom layer, while the field layer is
dominated by sedges. among which Cares lasiocarpa, Curex dioica and E. r,aginatum are most
common. The flarks have a much more sparse vegetation. Occasionally they have a thin bottom layer of
mosses. while Carex limosa. Carex magellanica,
Carex rostratu, Eriophorum angust(folium and Equisetum jhwiatile
occur
in the field layer. At
Bjommyran,
peat was collected at a flark, a string
and a site with a visible surface flow of water.
2.2. Peat collection and sample preparation
Peat profiles (down to about 50 cm depth at
Stor-Amyran and about 70 cm depth at Bjommyran)
were withdrawn from the ground with a peat auger.
Stor-Amyran was sampled in June 1991 and again in
September the same year, while Bjijmmyran
was
sampled in September 1992. The peat profiles were
cut and four 5-cm intervals distributed over the
profiles were put into separate plastic bags and
brought to the laboratory. The samples were stored
cool (IO- 15°C) for one night. The next morning, the
peat samples were cut with scissors and manually
mixed. Duplicate subsamples
were transferred to
flasks for the assay of potential methane oxidation
(described below). The remaining peat was frozen at
- 20°C and subsequently used for phospholipid analysis.
2.3. Phospholipid
analyses
While still frozen, the peat samples were partly
homogenized with a knife, and 2 g was transferred to
I. Sundh et al. / FEMS Microbiology
a glass tube with a teflon-lined
screw cap. Total
lipids were extracted using a modified one-phase
Bligh and Dyer extraction procedure [ 191. After addition of the extraction mixture to the tubes, the peat
was finely cut with a Polytron mixer. The total lipids
fraction was further fractionated by silicic acid chromatography [20] into neutral lipids (not retained by
the silicic acid column), glycolipids
(eluted with
acetone) and polar lipids, including the phospholipids (eluted with methanol). The polar lipid fraction
was dried under a stream of nitrogen and stored at
- 20°C until methanolysis.
The polar lipids were trans-esterilied
using a mild
alkaline methanolysis [21]. The methyl ester of nonadecanoic acid (19:O) (2.0 pg, = 6.4 nmol) was
added as an internal standard before methanolysis.
The fatty acid methyl ester preparations were stored
dry in glass tubes with teflon-lined
screw caps at
- 20°C until they were further processed.
Monounsaturated
fatty acids with 08 unsaturation
separate poorly from the corresponding
w7 and w9
isomers when analyzed by gas chromatography.
To
quantify the w8 isomers, methyl sulphide adducts of
the monounsaturated
fatty acids were prepared using
derivatization
of the double bonds with dimethyl
disulphide (DMDS) [22,23]. The derivatized samples
were then analyzed by gas chromatography/mass
spectrometry. The efficiency of the DMDS derivatization procedure was checked by analyzing derivatized and non-derivatized
standard PLFAs (16: 1 w7
and 18: 1~7) using gas chromatography.
The efficiency of conversion
was 90% (SD = 1%) for
16: 1 w7 and 69% (SD = 8%) for 18: I ~7. Concentrations of the w8 PLFAs in the peat samples were
corrected accordingly.
GC-MS analyses were performed on a HewlettPackard 5890A gas chromatograph connected to an
HP 5970B mass-selective detector (Hewlett-Packard,
Palo Alto. CA). The DMDS-derivatized
PLFAs were
separated on a DB5 fused silica capillary column (30
m X 0.25 mm, J&W Scientific, Folsom, CA). Samples of 1 ~1 were injected with an HP 7673A
autosampler in splitless mode at 50°C. The initial
temperature was held for 3 min and then raised from
50 to 160°C at 10°C per min, then at 4°C per min to
300°C. The linear flow velocity was 31 cm/s, injector temp 250°C MSD transfer line 3OO”C, and electron energy at ionization
70 eV. In the electron
Ecology 18 (1995) 103-I 12
105
impact mass spectra of DMDS adducts of monounsaturated fatty acids, cleavage of the fatty acid chain
between the -S-CH, moieties produces w- and Afragments with intense peaks diagnostic of the original position of the double bond. The MSD was set
up for selected ion monitoring using sets of ions
representing
the molecular ion (M + ) and the oand A-fragments of the DMDS adducts [23]. Quantification of l6:l w8 and l8:108 was done by monitoring the ions of the A-fragments at m/z 203 and
23 1, respectively. The M + ions at m/z
362 and
390, as well as the ions of the w-fragments at m/z
159. were used as qualifying
ions. Adducts were
identified by comparing their retention times with
data from standards obtained from Larodan AB
(Malmii. Sweden). The amounts of fragments specific for 16: 108 and 18: 1 w8 were quantified against
the internal standard (methyl ester of 19:O). The
coefficient of variation for replicate runs on the
GC-MS was 6%.
2.4. Calculation
bacteria
of cell numbers
of methanotrophic
Based on the literature, we first calculated that the
contribution
of 08 PLFAs is 33% (total reported
range 14-41 %IG)
of the total PLFAs in type I methanatrophic bacteria and 49% (total reported range 1968%) of the PLFAs in type II methanotrophic bacteria, respectively
[IO, 12.13,15,24]. These estimates
are averages of the published values.
Concentrations
of the two fatty acids were then
converted to cellular dry weight assuming that the
methanotrophic
bacteria contain 100 pmol PLFA
per g dry weight (d.w.) [10,25]. Since we cannot say
for certain which methanotrophs were present in the
peat, we used Methylomonas spp. and Methylosinus
trichosporium
as ‘type organisms’ for type I and
type II methanotrophs,
respectively.
Based on reported size ranges for these organisms [26-281 we
then estimated the volumes of type I and type II
methanotrophs
in the peat to 0.7 and 2.0 pm3,
respectively. These volumes are quite uncertain. For
example, based on the range in cell size for Methylomonas spp. reported by Whittenbury
and Krieg
[27], the range in cell volume is 0.16 to 2.9 pm’.
Next, the allometric expression of Norland et al.
[29] was used to calculate the percentage of cellular
106
1. Sundh et al. / FEMS Microbiology
Ecology
18 f 1995) 103-112
60
dry matter: 0.17 pg pm-” for a 0.7 pm” cell and
0.15 pg pm-”
for a 2.0 pm’ cell. We used this
expression without modification, since it is based on
analyses of a very large number of bacterial strains.
T
1
0
r’z0.63
p=0.0001
2.5. Potential methane oxidation
Duplicate 5- or 10-g samples of the homogenized
peat were transferred to 130-ml glass flasks with
screw caps equipped with butyl rubber stoppers.
Deionized water (20 or 25 ml) was added, and the
flasks were evacuated and refilled with air in three
cycles. Methane was added to a final concentration
of 0.3% (v/v> in the gas phase. The flasks were
incubated at 24°C on a rotary shaker and the methane
concentration in the gas phase was monitored for ca.
15 h. More details on the potential oxidation measurements have been given earlier [5,16].
The maximal methane oxidation rate per bacterial
biomass unit (V,,,)
in the peat was calculated by
dividing
the potential
oxidation
rate (nmole
min-’ g- ’ of wet peat) by the total dry weight of
methanotrophic
bacteria (mg g-’ of wet peat), as
estimated from the PLFA concentrations.
3. Results
Concentrations
in the peat of the PLFAs 16: 1 w8
and 18: 1 w8 ranged between 0.0 and 73 pmol g- ’ of
wet peat and 1.0 and 490 pmol g-’ of wet peat,
respectively. Pooling all the data (from both mires,
Potential
0
Stor-Amyran
Bjiirnmyran
Habitat
String
Mudbottom
Hollow
Minerotrophic
Hummock
Flark
String
‘Running’ water
min.’ g” wet peat)
1
2
Potential
(nmole
3
4
5
CH, oxidation
min”
g” wet peat)
Fig. 1. Relationships between concentrations of the specific PLFAs
and potential methane oxidation in peat. The regressions include
data from both mires, all sites and all depths 01 = 53).
all sites and all depths) resulted in significant correlations between potential methane oxidation and the
concentration of each PLFA (Fig. I ). The regressions
indicate a direct proportionality,
i.e. a doubling of
potential oxidation corresponds to a doubling of the
PLFA concentrations.
Concentrations
of the two
Table 1
Potential methane oxidation. total numbers of methanotrophic
bacteria, and average
communities; potential oxidation and cell numbers are expressed per g of wet peat
Mire
CH, oxidation
(nmole
biomass-specific
V,,,
Pot. oxidation a
nmol min- ’ g- ’
Cell numbers
millions g- ’
v Ill&Xh
pmol min-
0.07-4.1
0.00-2.7
0.00-0.3
0.00-3.2
0.02-I .5
0.00-4.3
o.oo- I .3
0.00-3.7
0.9-5 I
0.3-18
0.5- 1.9
1.5-19
0.5-I I
0.8-24
1.9-21
0.5-22
0.74
0.63
0.74
0.62
0.60
0.42
0.23
0.47
i The oxidation dataDhave previously been presented in another form by Sundh et al. 15,161
Values from Stor-Amyran are means of the June and September samples.
in peat from different
’ mg- ’ d.w. of cells
mire
1. Sundh
et ul. / FEMS
Microbiology
Ecology
Potential CH4 oxidation
4
6
2
f
10
3
/
/
/
40 I!T.l
0
0
107
O0
0,2
20
/
d;
,‘I
30 R
B
f
5
10
15
2
4
6
Cell numbers of methanotrophic
0,3
-“:y
10
0
/’
/
30
I2
(nmole min-1 g-1 wet peat)
1
o”
20
C199.5)103-I
and 5 1 X lo6 cells g- ’ of wet peat (Table 1). Highest numbers were found at the string site at StorAmyran, whereas numbers at all depths were relatively low at the wet hollow at Stor-Amyran.
Fig. 2 shows one example from each site GtorAmyran, September) of the depth distributions
of
potential methane oxidation and the numbers of type
I and type II methanotrophic bacteria. In most cases,
cell numbers closely followed the distribution
of
potential oxidation. Thus, both the highest cell numbers and the maximum of potential methane oxidation occurred close to the level of the water table.
The overall V,,, average, obtained by pooling of
all individual
peat samples,
was 0.60 prnol
PLFAs were strongly correlated with each other
(r’ = 0.67, P = 0.0001); i.e. whenever conditions in
the peat are favourable for aerobic methane oxidation, both type I and type II methanotrophic bacteria
are present.
The PLFA concentrations
indicated
a lower
biomass of type I (0.00-2.2
pg d.w. g-’ of wet
peat) than of type II methanotrophic
bacteria (0.029.7 /Lg d.w. g-’ of wet peat). Assuming that type I
cells are smaller, however, numbers of type I and
type II methanotrophic
bacteria within a particular
peat sample were similar. Comparing all sites and
depths, the total number (sum of type I and type II>
of methanotrophic bacteria ranged between 0.3 X lo6
2
18
4o0
,d/,
0.2
0,4
.cI
0,6
0.6
*
Potential CH4 oxidation
-
Cell numbers (type I)
- Q
Cell numbers (type II)
bacteria ( x 106 g-1 wet peat)
Fig. 2. Depth distributions of potential methane oxidation and cell numbers of type I and type II methanotrophic
bacteria in peat profiles.
A-E from Star-Amyran,
F-H from BjGrnmyran. A = string, B = mud-bottom.
C = hollow, D = minerotrophic
area, E = hummock.
F = flark, G = string and H = ‘running’ water. Note between-site differences in scales for both potential methane oxidation and cell
numbers of methanotrophic
bacteria. Horizontal lines show the depths of the water table at the time of sampling.
min-’ mg-’
d.w. of cells. The site-specific
V,,,
average (i.e. including all four depths) ranged from
0.23 pm01 min-’ mgg’ d.w. at the string community at Bjiimmyran to 0.74 pmol min- ’ rng-’ d.w.
at the string at Stor-Amyran (Table 1).
4. Discussion
4. I. PLFAs As biomarkers f<jr methanotrophic
ria
bacte-
If a compound (or group of compounds) is used
as a biomarker for specific microorganisms
in complex environmental
samples, several prerequisites
should be fulfilled [30]: (1) The compound must be
possible to extract and analyze with sufficient precision. (2) It must be present in a fairly uniform
concentration
within cells. (3) It should be sufficiently specific for the organisms to be quantified.
(4) It should be rapidly degraded in senescent and
dying ceils.
Several studies have shown that the first of these
prerequisites
(analytical
precision) is not a major
problem in the case of PLFAs. For example. when
bacteria (or pure bacterial phospholipids) are added
to soils or sediments the PLFAs can be quantitatively
extracted [ 19.3 l-331.
The second prerequisite (uniform concentration
within cells) is fairly well met by PLFAs. Thus, the
total PLFA content of eubacteria fluctuates around
ca. 100 pmol PLFA gg’ d.w. of cells [25], and
reported contributions of 16: 1 w8 and 18: 1 w8 to the
total cellular PLFA content are within a relatively
narrow range [ 10,12.13.15,24]. Variation in the total
PLFA content and in the contributions
of the w8
PLFAs are obviously related to species, but also to
the fact that PLFA composition may change due to
changes in environmental
conditions [34-361. However, as recently pointed out by Haack et al. [37],
existing data are not sufficient for a general evaluation of the impact of the environment
on PLFA
content and composition in microorganisms.
Another
matter of concern is that the two 08 PLFAs do not
occur in all isolates of methanotrophic bacteria. Thus,
16: 1 w8 has been encountered in some. but not all.
species of the genus Methylomonas.
Similarly,
18:1 w8 occurs in several, but not all, known genera
of type II methanotrophs
[ 12,241. Clearly, the fact
that these two fatty acids do not occur in all isolates
of methanotrophic
bacteria indicates that estimates
based on analysis of these PLFAs may underestimate
the total methanotrophic biomass.
Regarding the third prerequisite (the degree of
specificity), the PLFA 18: I w8 appears to be unique
for type II methanotrophic bacteria, since it has yet
to be found in any other organism [ 151. On the other
hand, it has been shown that small amounts of the
PLFA 16: 1w8 may sometimes be synthesized from
externally
added
16:0 in cultures
of Bacillus
strains[38]. In our data. however, the proportionality
between potential methane oxidation and PLFA concentrations suggests that the 16:1w8 content in the
peat reflects biomass of type I methanotrophic bacteria.
The final prerequisite, i.e. that compounds used as
biomarkers in environmental
samples must disappear
as the cells die, is well met by PLFAs. Thus, radioactively
labelled phospholipids
added to sediments or soils are rapidly hydrolyzed [ 19.39,40]. In
contrast to intact phospholipids, free fatty acids are
not retained by the silicic acid and therefore not
included in the final fatty acid extract. For our peat
samples, the proportionality
between concentrations
of the PLFAs and potential methane oxidation further implies that the extracted PLFAs occur in the
phospholipids of viable bacterial cells rather than in
dead material.
Even though the regressions of PLFA concentration against potential methane oxidation showed excellent statistical significance (P = 0.0001). and the
slopes of the regression lines were close to unity,
there was a fair amount of scatter. Several probable
sources for this can be identified: (1) Low analytical
precision of the GC-MS instrument. This is of minor
importance since replicate GC-MS runs had a coefficient of variation of only 6%. (2) Heterogenic distribution of the methanotrophs in the peat. Since larger
peat samples were used for the measurements
of
potential oxidation than for PLFA analysis, a patchy
distribution
of the methanotrophs
would introduce
scatter. Comparison of variance for replicate extractions and the regressions [41] suggests that variation
in replicate extractions explains at least 15% of the
variance in the regressions. (3) Differences in taxonomic diversity and composition (and hence differ-
I. Sundh et al. / FEMS Microbiology
ences in PLFA content and composition)
among
methanotrophic
communities
in different environments and depths. (4) Differences in PLFA content
and composition
among
similar
methanotrophic
communities
due to different environmental
conditions at different sites and depths. Due to lack of
data, it is presently not possible to evaluate the
importance of points 3 and 4 above.
4.2. Biomass and actbit?, of methanotrophic
in peat
bacteria
Our estimates of V,,, , with an overall average of
ca. 0.60 pmol methane consumed min- ’ mgg ’ dry
weight of organisms, agree well with V,,, estimates
for methanotrophic
bacteria. Thus, the V,,, of an
enrichment culture of methanotrophs
was ca. 0.60
pm01 min-’ mg- ’ d.w. of cells 1421, whereas V,,,
estimates for the type I methanotroph Methylococcus
range
from
0.19
to 0.75
pm01
capsulatus
min-l mg-’
d.w. of cells [43,44]. In contrast,
Jorgensen [45] reported that V,,, for the type II
methanotroph
Methylosinus
trichosporium
OB3b
was only 0.026 pmol mini’ mgg’ d.w.. It should be
kept in mind that our potential methane oxidation
rates are likely to be the sum of activities of several
species of methanotrophs exhibiting different biomass
specific V,,, values. Still, the close agreement between our V,,, estimates and those reported in the
literature for enrichments
and isolates of methanotrophs supports the assumption that concentrations
of the two fatty acids 16:l WI? and 18:l w8 provide
realistic estimates of the biomass and cell numbers
of methanotrophic
bacteria in this type of peatland,
and perhaps in other environments
as well.
To our knowledge,
only two previous studies
have reported cell numbers of methanotrophic bacteria in Sphagnum-dominated
peatlands [46,47]. Using
the most probable number (MPN) method, Williams
and Crawford [46] found 1.0 to 4.3 X 10’ cells ml- ’
of filtered peat pore water, which is three orders of
magnitude lower than our numbers. This large difference may reflect actual differences between the peatlands. However. we have found that the potential
methane oxidation activity in filtered peat pore water
is much lower than that of whole peat samples (data
not shown), indicating
that most of the methanotrophs in peat are attached to particles. Further-
Ecolog.~ 18 (1995) 103-112
109
more, it has been found that large numbers of
methanotrophic
bacteria in the surface water of waterlogged tundra environments can be attached to the
surfaces of particles of decaying organic matter [47].
The cell numbers reported by Williams and Crawford [46] are thus most likely underestimates because
a large fraction of the methanotrophs were removed
when the water was filtered. Another possibility is
that the MPN method per se gave an underestimation
since methods dependent on growth of the organisms
under laboratory conditions (i.e. plate counts and
MPN) often underestimate bacterial cell numbers in
natural environments
[48].
Vecherskaya et al. [47] used fluorescent antibodies to quantify methanotrophic bacteria in wet tundra
soils underlain
by permafrost. Their numbers of
methanotrophs were similar to ours and ranged from
0.1 to 22.9 X lo6 cells gg ’ of soil. In view of the
fact that their methods differed greatly from ours,
this similarity is remarkable. Vecherskaya et al. were
also able to distinguish between type I and type II
methanotrophic
bacteria. In most of their samples,
type I was approximately twice as abundant as type
II methanotrophs. Such a difference was not evident
in our samples. In this context, however, it should be
stressed that our cell numbers are strongly dependent
on the assumed cell volumes, which differed for type
I and type II methanotrophs.
In the peat profiles of this study, cell numbers of
methanotrophic bacteria were at a maximum close to
the depth of the water table. This is also the level
where both methane and oxygen is present, implying
that, as in other environments
having large methanatrophic populations,
the supplies of methane and
oxygen largely determine the biomass distribution of
methanotrophic bacteria [3,4,6].
The close coupling between the concentrations of
the two w8 PLFAs and the potential methane oxidation activity has several implications: (I) The investigated type of peatland harbours high numbers of
viable methanotrophic
bacteria, which immediately
oxidize methane at rates close to V,,, when conditions are favourable. (2) The coupling between potential oxidation activity and the concentrations
of
the two PLFAs suggests that representatives
of
‘classical’ type I or type II methanotrophic
bacteria
are prominent in the methanotrophic communities of
the peats. Possibly, methanotrophic
yeasts [49], am-
monia-oxidizing
bacteria able to metabolize methane
[50], or unknown methanotrophs,
are of minor importance. (3) Our finding that the concentrations
of
the w8 PLFAs were strongly correlated with each
other shows that whenever conditions in the peats
are favourable for aerobic methane oxidation, type I
and type II methanotrophic bacteria occur simultaneously.
In conclusion, we find that the points addressed
above support our conclusion that concentrations
of
the PLFAs 16:l w8 and 18:l w8 give reliable estimates of the biomass and cell numbers of methanatrophic bacteria in the peat samples. This was further substantiated by the observation that cell numbers derived from concentrations
of PLFAs agreed
well with corresponding estimates based on the fluorescence antibody technique [47].
Acknowledgements
We are grateful to Eva Lindstrom for skilful
preparation of derivatized phospholipid
fatty acids
tut of frozen peat. We also thank Anders Tunlid and
Asa Frostegird
at the Department
of Ecology at
Lund University,
Lund, Sweden, for sharing their
great knowledge of methods for lipid isolation and
quantification.
This work was financially supported
by the Swedish Council for Forestry and Agricultural
Research (contract 33.04001, the Swedish National
Board for Industrial and Technological Development
(contract C 216 097-2), and the Swedish Environmental Protection Board (contract 27204).
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