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Applied and Environmental Microbiology, June 1999, p. 2679-2684, Vol. 65, No. 6
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Nitrous Oxide Production and Methane Oxidation by
Different Ammonia-Oxidizing Bacteria
Qing-Qiao
Jiang1 and
Lars R.
Bakken2,*
Department of Biotechnological
Sciences1 and Department of Soil and
Water Sciences,2 Agricultural University of
Norway, 1432 Aas, Norway
Received 28 September 1998/Accepted 29 March 1999
 |
ABSTRACT |
Ammonia-oxidizing bacteria (AOB) are thought to contribute
significantly to N2O production and methane oxidation in
soils. Most of our knowledge derives from experiments with
Nitrosomonas europaea, which appears to be of minor
importance in most soils compared to Nitrosospira spp. We
have conducted a comparative study of levels of aerobic N2O
production in six phylogenetically different Nitrosospira
strains newly isolated from soils and in two N. europaea
and Nitrosospira multiformis type strains. The fraction of
oxidized ammonium released as N2O during aerobic growth was
remarkably constant (0.07 to 0.1%) for all the
Nitrosospira strains, irrespective of the substrate supply
(urea versus ammonium), the pH, or substrate limitation. N. europaea and Nitrosospira multiformis released
similar fractions of N2O when they were supplied with ample
amounts of substrates, but the fractions rose sharply (to 1 to 5%)
when they were restricted by a low pH or substrate limitation.
Phosphate buffer (versus HEPES) doubled the N2O release for
all types of AOB. No detectable oxidation of atmospheric methane was
detected. Calculations based on detection limits as well as data in the
literature on CH4 oxidation by AOB bacteria prove that none
of the tested strains contribute significantly to the oxidation of
atmospheric CH4 in soils.
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INTRODUCTION |
Nitrogen transformation in soil and
water is considered one of the main reasons for the increasing
N2O levels in the atmosphere (6), and
ammonia-oxidizing bacteria (AOB) appear to be important in this process
(7, 8, 10, 27-29). The immediate (but partial) inhibiting
effects of specific nitrification inhibitors on N2O production in soils (2, 5, 7, 11) demonstrate the direct role of nitrifier metabolism in producing N2O. AOB are also
thought to be involved in the oxidation of atmospheric methane by
aerobic soils, since methane is an alternative substrate for ammonia
monooxygenase (3, 19). Methane oxidation in soil and its
inhibition patterns may indicate that ammonia oxidizers are involved
directly or indirectly (1, 15, 37). The kinetics of methane
oxidation observed in type strains of AOB so far suggest a minor role,
however (1, 13, 20). Thus, unless significantly higher
oxidation rates are demonstrated in new isolates, their direct
participation appears insignificant.
Nitrosomonas europaea appears to produce N2O by
more than one mechanism. Moderate amounts are released under full
aeration, but the release increases sharply in response to oxygen
limitation (12, 24). Poth and Focht (23) showed
that N. europaea denitrified with
NO2
as the electron acceptor and that the
labelling pattern observed (with either
15NH4+ or
15NO2) indicated that
N2O was primarily a product of
NO2 reduction, rather than a by-product of
NH3 oxidation. The presence of nitrite reductase in
N. europaea has been demonstrated in several investigations
(9, 22, 25, 26), and it is probably involved in the
production of N2O by this organism under oxygen-limiting conditions (17). Under full aeration, however, the
production of small amounts of N2O by N. europaea seems to be a direct by-product of NH2OH
oxidation to NO2, as judged from the pattern
of inhibition and substrate dependency of the process (17).
This possibility implies that the enzyme hydroxylamine oxidoreductase
is directly involved in N2O production, since it catalyzes
complete NH2OH oxidation to NO2
without any free intermediates (38). The possible
involvement of hydroxylamine oxidoreductase in the release of
N2O hypothetically implies that any external factor of
importance for this periplasmic enzyme may affect the
N2O/NO2 product ratio.
Experiments with whole soil communities have suggested that acidity as
such results in a high product ratio (21).
Nitrosomonas is the most commonly used genus in laboratory
studies of AOB, but Nitrosospira seems to be the most
dominant species in natural environments (14, 32). On this
basis, we decided to do a comparative investigation of levels of
N2O production in AOB, using two N. europaea and
Nitrosospira multiformis (see below) type strains and six
Nitrosospira strains which have recently been isolated from
various terrestrial environments (18, 33, 34). The
experiment allowed CH4 concentrations to be measured along
with N2O, thus allowing us to check whether any of the
cultures were able to oxidize significant amounts of methane at
atmospheric concentrations. This approach is insensitive compared to
tracer (14C) methods commonly used for measuring
CH4 oxidation by AOB (3). However, calculations
showed that if a culture had anywhere near the oxidation rates
necessary for AOB to play a role in soil, we would easily detect and
measure its oxidation in our experiments.
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MATERIALS AND METHODS |
Cultures.
Six strains of AOB isolated from acidic forest
soils (strains III2, III7, and L115), acid sandy loam from Zambia
(strain AF), neutral clay loam soil (strain 40K1), and wastewater from
a treatment facility (strain B6) were used. These strains were
identified as Nitrosospira spp. based on morphology and
their 16S ribosomal DNA (rDNA) sequences (18, 33, 34). One
N. europaea culture (provided by J. Prosser, University of
Aberdeen, Aberdeen, Scotland) and the Nitrosolobus
multiformis type culture ATCC 25196 (American Type Culture
Collection, Rockville, Md.) were also used. We assume that a new name
for the ATCC 25196 strain will be Nitrosospira multiformis
because of the 16S rDNA-based consensus to merge the genera
Nitrosospira, Nitrosolobus, and
Nitrosovibrio into one genus, Nitrosospira
(33, 34). Herein we use the name Nitrosospira multiformis.
Medium.
The basic medium used was a liquid mineral (LM)
medium containing (per liter) KH2PO4 (0.2 g),
CaCl2 · 2H2O (0.02 g),
MgSO4 · 7H2O (0.04 g), FeNaEDTA (3.8 mg), phenol red (0.1 mg), NaMoO4 · 2H2O
(0.1 mg), MnCl2 (0.2 mg), CoCl2 · 6H2O (0.002 mg), ZnSO4 · 7H2O (0.1 mg), and CuSO4 · 5H2O (0.02 mg). The N source and the buffering of the LM
medium for each experiment are reported below. The N source was added
from stock solutions containing either 1.0 M
(NH4)2SO4 or 1.0 M urea (analysis
grade; Merck). The medium and the stock solutions were sterilized by
autoclaving at 121°C for 20 min.
Culturing and harvesting of cells.
The cell inoculum for the
experiment was grown in 150-ml batches of medium in 500-ml E flasks.
The medium was the LM medium with 3.8 mM
(NH4)2SO4 and 15 mM HEPES sodium
salt (art. 15231; Merck); its pH was adjusted to 7.5. The flasks were
incubated on a reciprocal shaker (100 rpm) at 21°C in the dark, and
the cells were harvested by filtration with a 0.2-µm-pore-size
polycarbonate membrane (Nuclepore, Costar Europe Ltd., Badhoevedorp,
The Netherlands). For harvesting of about 300 ml of culture, three to
four membranes had to be used due to clogging. The cells were washed
off the filters and dispersed in about 15 ml of substrate-free LM
medium. The dispersion was obtained by pumping the cells through a
0.5-mm-diameter needle a dozen times. The cell density was determined
by fluorescence microscopy after staining the cells with acridine
orange. The inoculum was kept at 21°C in the dark and used within
3 h.
Incubation experiments for determination of N2O
production and CH4 oxidation.
The incubation
experiments were designed to investigate N2O production as
well as CH4 consumption during growth on
NH4+ or urea. In the first experiment, the
medium [LM medium supplemented with 3.8 mM
(NH4)2SO4 or urea] was buffered
with 15 mM HEPES. Since the cultures were to be prevented from being
exposed to atmospheric CO2, the medium contained 0.84 mM
sodium carbonate to avoid CO2 limitation of growth
(assuming that 0.11 mol of C is assimilated per mol of
NH4+ oxidized [4], this
quantity should be more than sufficient). The pH was regulated to 7.5. Batches of 20 ml of medium were placed in 120-ml serum flasks and
inoculated to reach an initial cell density of 1 × 107 to 2 × 107 cells ml1.
The flasks were then sealed with Teflon-faced silicone septa (20-ST3;
Chromacol Ltd., Herts, United Kingdom) and incubated at 25°C in the
dark (no methane was added). The cultures were stirred with a 2-cm-long
magnetic bar in each flask, operated with pulsed stirring (60 s of
stirring at 300 rpm and 15-s pauses) with a Telemodul 20P (Variomag,
Munich, Germany) equipped with a 15-flask-position magnetic plate
(Variomag model no. HP15) submersed in a thermostated water bath. The
samplings were done periodically during the incubation. Gas samples
were taken directly from the serum flasks into the injection loop of
the gas chromatograph via a peristaltic pump system equipped with a
side port needle to penetrate the silicone septa. Liquid samples for
NH4+ and NO2 analysis
were taken with sterile syringes (through the silicone septa) and
frozen to 
20°C until analysis.
The results of the first experiment suggested that some of the cultures
increased the relative production of N2O as the culture reached a critically low pH. The second experiment was designed to test
if this pattern was reproducible and to increase the resolution of the
experiment by changing the buffer and monitoring the pH along with the
production of NO2 and N2O. To
prolong the period of activity in the critical pH range (pH 5.3 to
6.4), 20 mM phosphate buffer was used instead of HEPES (based on the
titration curves for the two buffers for this pH range). The buffer
consisted of NaH2PO4 and
Na2HPO4 in a 31:9 ratio, giving an initial pH
of 6.4 in the LM medium (at 20 mM phosphate). The cells for the
inoculum were grown and harvested as in the previous experiment and
resuspended in substrate-free 20 mM phosphate-buffered LM medium. The
serum flasks were filled with 40 ml of 20 mM phosphate-buffered LM
medium containing 7.6 mM NH4+ and sealed with
Teflon-faced silicone septa (type 20-ST3; Chromacol Ltd.), and 0.5 ml
of CO2 was added to ensure the presence of C for growth
(the medium contained no carbonate). The amount of oxygen in the
remaining 80 ml of gas headspace was about 0.7 mmol per flask, which is
about twice the molar amount of NH4+ per flask.
Thus, if complete oxidation of all the NH4+
occurs (1.5 mol of O2 per mol of NH3), the
cultures would approach oxygen-limiting conditions. However, the
buffering capacity of the medium was deliberately made so weak that
acid inhibition of growth would occur before 50% of the
NH3 was oxidized.
Chemical analyses.
The gas concentrations were determined
with a gas chromatograph equipped with three detectors (thermal
conductivity, flame ionization, and electron capture detectors)
allowing sensitive and rapid measurement of N2O,
CH4, and CO2 with a single injection (30). The sampling system extracts 3 ml for each
determination. The pressure reduction resulting from repeated samplings
from the same bottle was accounted for when we calculated the amounts of N2O and CH4 (reported amounts are the values
that would be reached if sampling did not take place).
A flow injection system (FIA star; Tecator AB, Höganäs,
Sweden) was used to analyze NH
4+ and
NO
2. Nitrite was determined by color reaction
from Griess-Ilosvay
reagents I and II and was measured at a wavelength
of 540 nm.
Ammonium was determined by merging a sample with NaOH to
form
gaseous NH
3, which passed through a Teflon membrane to
a pH indicator
mixture (Application Sub Note ASN 151-01/92; Tecator).
pH was
measured with a 6-mm-diameter pH electrode (ROSS Semi-micro
Combination
pH electrode, model 8103; ATI Orion, Boston, Mass.), which
allows
measurement in small sample
volumes.
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RESULTS |
Figures 1 and
2 show the accumulations of
N2O (in micrograms of N per flask) and
NO2 (in milligrams of N per flask) when cells
were grown in HEPES-buffered medium supplemented with
NH4+ and urea, respectively. The III7 and AF
cultures failed to grow in this experiment (data not shown). For some
unknown reason, strain B6 did not show significant activity in the urea
medium (Fig. 2), despite its proven urease activity in a previous
experiment (unpublished). Slight production of
NO2 and N2O was found in N. europaea and Nitrosospira multiformis cultures. The
NO2 concentrations in the stationary phase
were between 0.5 and 1 mg of NO2-N per flask, which is 25 to 50% of the total NH4+-N in the medium
(i.e., 2.1 mg of N per flask). In contrast, the urea-grown
urease-positive strains (40K1 and III2) accumulated around 1.5 mg of
NO2-N, i.e., around 75% of the added urea N
(Fig. 2). The final pH values for each culture are shown in Table
1.
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FIG. 1.
Accumulation of N2O (in micrograms of N per
flask) and NO2 (in milligrams of N per flask)
during growth with ammonium as the substrate in the HEPES-buffered
medium (initial pH, 7.5).
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FIG. 2.
Accumulation of N2O (in micrograms of N per
flask) and NO2 (in milligrams of N per flask)
during growth with urea as the substrate in the HEPES-buffered medium
(initial pH, 7.5).
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The accumulation of N2O-N was very close to 0.1% of the
NO2-N accumulated for all the strains of
Nitrosospira through the whole growth cycle when the strains
were grown on both NH4+ and urea (Fig.
3). The same was true for the early phase
of NH4+-grown N. europaea and
Nitrosospira multiformis, but in these cultures the
accumulation of N2O continued after the
NO2 concentrations reached a stable level.
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FIG. 3.
Accumulated N2O-N as percentages of
accumulated NO2-N in cells grown in the
HEPES-buffered medium with ammonium (A) and urea (B). N.eur., N. europaea; N.multif., Nitrosospira multiformis.
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Methane concentrations were measured through the whole incubation
experiment, but the level (2.1 ml liter1 [ppmv])
remained practically constant (within 2.10 ± 0.02 ppmv [mean ± standard deviation]) throughout the whole incubation
(data not shown). Approximate upper confidence limits for possible
CH4 oxidation in the cultures was thus around 0.1 nmol of
CH4 per flask.
The results of the second experiment are shown in Fig.
4
to 7. In this experiment, the medium
(NH4+ only) was buffered with 20 mM phosphate
to enhance the resolution of the experiment in the desired pH range and
pH was measured in every sample (Fig. 4). The
NO2 concentration stabilized at different
levels, reflecting the pH tolerance of each strain. The stable levels
reached for NO2 accumulation did not exceed
0.5 mg of N per flask, which is only 12% of the
NH4+-N in the medium.
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FIG. 4.
Measured pH in the cultures growing in the
phosphate-buffered medium. N.multif., Nitrosospira
multiformis; N.eur., N. europaea.
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FIG. 5.
Accumulation of NO2 (in
milligrams of N per flask) during growth in the phosphate-buffered
medium. N.multif., Nitrosospira multiformis; N.eur.,
N. europaea.
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FIG. 6.
Accumulation of N2O (in micrograms of N per
flask) during growth in the phosphate-buffered medium. N.eur., N. europaea.
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FIG. 7.
Yields of N2O-N as percentages of
NO2-N in the phosphate-buffered medium,
plotted against the measured pH. N.multif., Nitrosospira
multiformis; N.eur., N. europaea.
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The accumulation of N2O (Fig. 6) resembled that of
NO2 (Fig. 5), as was found in the first
experiment with HEPES-buffered media (Fig. 1 and 2). The
N2O production by Nitrosospira multiformis culture (data not shown in Fig. 6) showed a pattern similar to those of
the other cultures, but the production was much higher (final amount, 3 µg per flask). The relative production of N2O (as a
percentage of NO2 produced) and its
relationship with pH in the medium are illustrated in Fig. 7, where
accumulated N2O as a percentage of
NO2 is plotted against the measured pH. For
the Nitrosospira cultures, the level of N2O
increased gradually through the first part of the growth period but
reached more or less stable levels between 0.2 and 0.3%. In contrast,
N. europaea and Nitrosospira multiformis continued to higher levels (0.4 and 0.5%, respectively). Thus, the
experiment with phosphate buffer reproduced the pattern of the previous
experiment as far as the contrast between the Nitrosospira strains and the two type strains is concerned, but the relative levels
of N2O production of the Nitrosospira strains
were higher and more variable in phosphate-buffered medium than in
HEPES-buffered medium.
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DISCUSSION |
Production of N2O.
The first experiment with cells
grown in HEPES-buffered medium with an initial pH of 7.5 (Fig. 1 to 3)
demonstrated that all cultures, including N. europaea
and Nitrosospira multiformis, had surprisingly similar
N2O/NO2 product ratios
during unrestricted growth. For the urease-positive Nitrosospira strains, the product ratios with urea as the
substrate were practically identical to those with
NH4+ as the substrate, and the rankings of the
cultures (III2 > 40K1 > L115) were identical for the two
substrates (Fig. 3). The product ratios were in the lower range of
those observed in other ammonia-oxidizing cultures during aerobic
unrestricted growth (12, 17).
In response to unfavorable conditions, however, the two type strains
(
N. europaea and
Nitrosospira multiformis)
differed significantly
from the
Nitrosospira strains. When
the oxidation of NH
3 to NO
2
became acid limited (Fig.
1), the two type strains continued
to produce
N
2O for some time, which resulted in significantly
higher
product ratios towards the end of the incubation (Fig.
3A). The
relative levels of production of N
2O over
NO
2, if calculated for single time increments
in the transition between
active phase and stationary phase (50 to
100 h for
N. europaea and 150 to 200 h for
Nitrosospira multiformis), were in the range
of 1 to 5%
(N
2O-N production as a percentage of
NO
2-N production). The type strains also
showed high product ratios
when they were grown in the urea medium
(Fig.
3B). The production
of NO
2 by the
urease-negative type strains in the urea medium was due
to traces of
NH
4+ present in the medium (some hydrolysis of
urea occurred during
the autoclaving). This result implies that the
cultures were active
but severely starved in this medium. Hence,
limitation of the
supply of NH
4+ seems to
affect the product ratio in the same way that acidity
did. We may be
reporting the same phenomenon (starvation), however,
since the pH
controls the concentration of NH
3, which is the substrate
for ammonia monooxygenase (
18,
38). The half-saturation
constant
(
Km) for some of the
Nitrosospira strains (AF, L115, B6, and 40K1)
ranged from 6 to 11 µM NH
3, which is comparable to the levels
of
affinity in the type strains (
18). Thus, differences in
affinity
hardly explain the differences in the levels of
N
2O production
between the type strains and the
Nitrosospira strains. The apparent
Kms of indigenous AOB in forest soils have been
found to be 2
to 3 orders of magnitude lower (10 to 15 nM
NH
3 [
31]) than those
of all cultured
organisms, suggesting that certain soils harbor
AOB essentially
different from those cultivated so far, including
the
Nitrosospira strains used in this
study.
The second experiment was designed to improve determination of the
regulating effects of pH and substrate supply, by using
an initial pH
of 6.4 in a weakly (20 mM) phosphate-buffered medium.
The measured pH
demonstrated that this aim was largely reached,
in terms of several
measurements of both NO
2 and N
2O
production as the cultures approached their characteristic
critical
pHs. These limits agree reasonably well with direct observations
of
each strain's pH tolerance (strains were exposed to preset
pH values
of 5 to 8.5; cf. with the work of Jiang and Bakken
[
18]).
Strain AF, however, is unlikely to be near
its critical pH level
in the present experiment, since it was found to
be at least as
acid tolerant as L115 and B6 (
18). The
explanation may be low
viability in the
inoculum.
The production of N
2O as a percentage of
NO
2 in the phosphate-buffered medium (Fig.
7)
contrasts with the results obtained
with the HEPES-buffered medium in
several ways. The product ratio
increased gradually through the first
50 to 100 h of the incubation,
both for the
Nitrosospira strains and for the two type strains.
The
Nitrosospira strains which reached a stationary-phase level
of production (L115, B6, III7, and III2) also stabilized the product
ratios at levels around 0.2% (Fig.
7), which is at least twice
the
level found in the HEPES-buffered medium (Fig.
3). The ranking
of
relevant
Nitrosospira strains (III2, 40K1, and L115) in the
phosphate buffer (40K1 > III2 = L115, final ratios) (Fig.
7)
was
different from that in the HEPES buffer (III2 > 40K1 > L115).
The only pattern in experiment 1 (HEPES-buffered medium) which
was really reproduced in experiment 2 was the ability of
N. europaea and
Nitrosospira multiformis to continue the
production of N
2O
longer than the production of
NO
2 when it was confronted with acid
limitation of activity, resulting
in higher final relative
N
2O values than those of the
Nitrosospira strains (Fig.
7).
The effect of buffer systems on N
2O production by
N. europaea was investigated by Hynes and Knowles (
17),
and they found
that a HEPES-buffered medium (pH 7.5) resulted in much
lower relative
N
2O production than a phosphate-buffered
medium at the same pH,
which is in agreement with our results. In
contrast, they observed
that two other organic buffer
systems [
N-tris(hydroxymethyl)methylglycine (pH 8) and
tris(hydroxymethyl)aminomethane
(pH 8.5)] resulted in relative
N
2O production much higher
than that of the phosphate buffer systems.
It seems clear that the
production of N
2O, which probably occurs
in the periplasm
(due to the location of hydroxylamine oxidoreductase
in this
compartment), can be strongly affected by a number of
external factors
other than pH and oxygen concentrations. This
conclusion implies that
one should not extrapolate the present
values for N
2O
production to populations in natural environments
without
reservations.
The purpose of the present investigation was primarily to investigate
whether consistent differences exist between AOB in
their abilities to
produce N
2O. It seems reasonable to conclude
that no such
consistent differences exist among the recently isolated
strains of
Nitrosospira. On the other hand, all the
Nitrosospira strains produced consistently less relative
amounts of N
2O than
the two type strains,
N. europaea and
Nitrosospira multiformis.
The relatively
high N
2O production by the two type strains occurred
only
during restricted activity, either due to acidity (Fig.
3A
and
7) or
due to low substrate supplies (Fig.
3B). A correlation
between acidity
and the relative N
2O production rate was found
for the
Nitrosospira strains as well, but it was weaker than for
the
two type strains and dependent on the buffer system
used.
Populations of AOB in the natural environment are likely to be
frequently confronted with starvation as well as critically
low pH
values (
35). The present investigation strongly suggests
that the ammonia oxidized by such starving and/or acid-limited
populations is likely to yield more N
2O than that released
by
an exponentially growing culture under optimal conditions. This
effect apparently depends on the composition of the AOB
community.
Oxidation of CH4.
In this study, no significant
oxidation of atmospheric CH4 was observed. This result may
be attributed primarily to the insensitivity of our approach. The
available data on methane oxidation by AOB have been obtained by using
radioactively labelled methane, which allows extremely low rates to be
determined. Hyman and Wood (16) found that CH4
oxidation by N. europaea had a Vmax
of 0.065 mmol of CH4 g (dry weight) of cells1
h1 and a Ks of 6.6 µM
CH4. In the present experiment, the CH4
concentration in the gas phase was around 2 ppmv, which means that the
CH4 concentration in the liquid medium was around 2.5 nM
(assuming equilibrium and with the solubility of CH4 being
calculated according to the method of Wilhelm et al.
[36]). If we use the Vmax and
Ks values reported above, we find that the
oxidation rate in our flasks should be 25 nmol g (dry weight) of
cells1 h1. Assuming a cell dry weight of
0.06 × 1012 g and an average cell density of 6 × 107 cells ml1 in the 40-ml medium per
flask in the second experiment, we obtain an oxidation rate of
approximately 5 pmol flask1 h1, which is
equivalent to 0.1% h1 (total methane in the flask was 6 nmol). Thus, if the organisms were actively oxidizing methane through
the entire incubation period of 320 h, the methane concentration
in the flasks should have been reduced by approximately 30%.
If ammonia oxidizers behaved according to the enzyme kinetics referred
to above, we would need 2 × 10
9 cells g (dry weight)
of soil
1 to account for the observed CH
4
oxidation in forest soils (1
to 3 ng of CH
4
g
1 day
1 [
30]). This cell
density is at least 3 orders of magnitude
higher than the density of
AOB found in soils. Thus, if AOB were
to contribute to the observed
CH
4 oxidation in aerobic soils,
there would have to exist
species that oxidize atmospheric CH
4 at a rate orders of
magnitudes faster than that observed to date.
If the
Nitrosospira strains had anywhere near such high rates
of
methane oxidation, we would be able to measure their activities
with
the present experimental design (i.e., without radioactive
tracers). We
did not and conclude that none of our strains are
likely to have
contributed significantly to the oxidation of atmospheric
CH
4 in the soil from which they were
isolated.
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ACKNOWLEDGMENTS |
This work was financed by the Norwegian Research Council, project
no. 114038/111, which is a part of the Nordic (NKJ) research program
Ecological Roles of Ammonia-Oxidizing Bacteria in Soils.
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FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Soil and Water Sciences, Agricultural University of Norway, P.O. Box 5028, 1432 Aas, Norway. Phone: 47 64948219. Fax: 47 64948211. E-mail:
lars.bakken{at}ijvf.nlh.no.
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1993.
Methane oxidation in temperate and subarctic forest soils: rates, vertical zonation, and responses to water and nitrogen.
Appl. Environ. Microbiol.
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Aulakh, M. S.,
D. A. Rennie, and E. A. Paul.
1984.
Acetylene and N-serve have effects upon N2O emissions from NH4+ and NO3 treated soils under aerobic and anaerobic conditions.
Soil Biol. Biochem.
16:351-356.
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| 3.
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Bedard, C., and R. Knowles.
1989.
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Abstract
Introduction
