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Applied and Environmental Microbiology, June 2000, p. 2636-2640, Vol. 66, No. 6
Department of
Microbiology,1 Department of Crop and
Soil Science,2 and The Laboratory for
N2 Fixation Research, Department of Botany and Plant
Pathology,3 Oregon State University, Corvallis,
Oregon 97331
Received 12 November 1999/Accepted 28 March 2000
Little information exists on the potential of
NH3-oxidizing bacteria to cooxidize halogenated
hydrocarbons in soil. A study was conducted to examine the cooxidation
of methyl bromide (MeBr) by an NH3-oxidizing bacterium,
Nitrosomonas europaea, under soil conditions. Soil and its
water content modified the availability of NH4+
and MeBr and influenced the relative rates of substrate
(NH3) and cosubstrate (MeBr) oxidations. These observations
highlight the complexity associated with characterizing soil
cooxidative activities when soil and water interact to differentially
affect substrate and cosubstrate availabilities.
In recent years considerable
research has been conducted to determine the soil physical and chemical
factors which control the fate of the agriculturally applied soil
fumigant methyl bromide (MeBr) (2, 3, 11, 25, 26, 29).
Although the potential of methanotrophic and NH3-oxidizing
bacteria to cooxidatively degrade MeBr has been known for some time
(12, 17, 20, 24), and facultatively methylotrophic soilborne
bacteria have been isolated that grow on MeBr as a C source (6,
15), only recently was it shown that soil bacteria act as a sink
for MeBr in situ (7, 15, 18, 22). At this time it is unclear
to what extent MeBr consumption in soil occurs by cooxidative rather
than energy-gaining metabolism and how soil factors might influence
microbiological MeBr transformation. Nitrosomonas europaea,
a chemolithotrophic NH3 oxidizer, carries out cooxidation
of a variety of halogenated and nonhalogenated hydrocarbons in the
presence of NH4+ through the activity of
ammonia monooxygenase (AMO) (12, 13, 19, 20, 21). Many
factors that influence NH3 oxidation in soil will
presumably influence transformation of alternate substrates. For
example, soil colloids are known to bind NH4+,
which might influence the ratio of substrate (NH3) to
cosubstrate during cooxidation and affect competition for the active
site of AMO. Hommes et al. (8) examined the ability of
N. europaea to oxidize NH4+ and
three cosubstrates, ethylene, chloroethane, and 1,1,1-trichloroethane, in vigorously aerated soil slurries. The influence of soil exchangeable acidity on solution pH and NH3-NH4+
equilibrium was the main factor affecting NH3 and
cosubstrate oxidation, whereas NH4+ adsorption
played a lesser role under slurry conditions. The influence of intact
soil on cooxidation by NH3 oxidizers could be more complex.
For example, in a study which examined the effect of soil water content
on nitrification, it was concluded that the primary negative effect on
nitrification of lowering soil water potential from saturation to Cell growth, experimental manipulations, and analyses.
Surface
samples of a Willamette silt loam (0 to 20 cm) were used, and the
properties are described elsewhere (14). The soil pH was
raised to approximately neutral (7.0 ± 0.2) by incubation for 3 days with 3 g Ca(OH)2 kg of soil
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Copyright © 2000, American Society for Microbiology. All rights reserved.
Effects of Soil and Water Content on Methyl Bromide Oxidation by
the Ammonia-Oxidizing Bacterium Nitrosomonas
europaea
ABSTRACT
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MPa was reduced diffusion of NH4+ and
NH3 to the sites of NH3-oxidizing activity
(23). Furthermore, it is not difficult to conceptualize that
MeBr oxidation might be sensitive to soil water content because of the
large differences in the diffusion coefficients of MeBr through water
and air (diffusion coefficients, 0.1037 cm2
s
1 for air and 1.35 × 105
cm2 s1 for water) (26). Recent
studies showed that relatively small changes in soil water content had
a profound effect on microbially mediated MeBr uptake by a forest soil
(7). The purpose of our research was to build upon earlier
studies (8) and to examine the characteristics of
NH3 and MeBr oxidations by N. europaea incubated
in intact soil held at different water contents.
1, air
dried, and sterilized with gamma irradiation (4 megarads) at the Oregon
State University Radiation Center. Batch cultures of N. europaea (ATCC 19718) were grown as described elsewhere (9). Late-exponential-phase cells (3 to 4 days) were
harvested by centrifugation (11,000 × g; 15 min),
washed twice in ice-cold buffer (50 mM potassium phosphate buffer, pH
7.2), and resuspended to a cell density of (30 ± 7) × 107 ml1 (80 ± 20 µg [dry weight] of
cells ml1).
1) and provide a total water
volume of 5 ml in 11 g of oven-dried soil. To conduct experiments
with saturated soil at WHC, the following mixture (prepared at 4°C
and kept in the dark on ice) was injected slowly through the septum of
each vial using a plastic syringe fitted with a 23-gauge needle: 25 or
100 mM NH4+ in 50 mM
K2HPO4 (pH 7.2), 2 ml; N. europaea
cell suspension with an optical density of 0.25, 1 ml; and 50 mM
K2HPO4 (pH 7.2), 1.4 ml. Vials without soil
were set up in a similar manner with an additional 0.6 ml of sterile
deionized water to provide the same total volume of water as in the
soil treatments. The final cell density of N. europaea was
equivalent to 6 × 107 cells ml of
water1 or 2.7 × 107 cells g of
soil1. To conduct experiments with unsaturated soil at
approximately two-thirds of WHC (300 g of H2O kg of
soil1), the same amounts of soil and MeBr were dispensed
into vials as described above. A 2.4-ml aliquot of water is required to
provide a total volume of 3 ml. The following mixture was injected into each vial: 25 or 100 mM NH4+ in 50 mM
K2HPO4 (pH 7.2), 1.2 ml; N. europaea
cell suspension with an optical density of 0.25, 0.6 ml; and 50 mM
K2HPO4 (pH 7.2), 0.6 ml. Vials without soil
were set up in a similar manner with an additional 0.6 ml of water. In
this case, we chose to keep the N. europaea cell density
constant on the basis of water; the density per gram of soil was lower
than in the saturated experiment (1.63 × 107 cells g
of soil1). Triplicate vials of each treatment were
incubated horizontally in the dark at 27°C with periodic rotation to
facilitate gas distribution. Agitation was avoided to prevent the
breakdown of soil structure, which would occur especially under
water-saturated conditions. Preliminary experiments were conducted to
determine if O2 limitation would occur in unshaken vials
and retard NO2 production and MeBr oxidation.
No differences were detected in rates of NO2
production assayed in unshaken vials containing either ambient or
supplemental levels of O2 (0.4 atm) at 10 or 40 mM
NH4+.
concentration determinations, 17 or
19 ml of ice-cold 2 M KCl (supplemented with 0.1 mM allyl thiourea, an
inhibitor of NH3 oxidation) was added to each vial
containing either saturated or unsaturated soil, respectively (2:1
liquid/soil ratio). Vials were shaken vigorously for 5 min and were
centrifuged to pellet the soil, and NO2
concentrations were determined in the supernatants (5). To measure the amount of NH4+ in soil solution
under saturated conditions, 4.4 ml of buffer containing either 50 or
200 µmol of NH4+ was added to 11.6 g of
air-dried soil in a centrifuge tube and was incubated for 1 h.
Following the incubation, soil was centrifuged and samples of
supernatant were recovered for analysis of
NH4+. To measure the amount of
NH4+ in solution under unsaturated conditions,
larger portions of soil (18.4 g of air-dried soil) were placed in 60-ml
syringes, and buffer (6 ml) containing 100 or 400 µmol of
NH4+ was dribbled over the soil. Samples were
incubated for 1 h and samples of soil solution were forced from
the soil by applying pressure with the plunger of a syringe. Samples
were centrifuged to pellet soil, and NH4+
concentrations were determined in supernatants after alkaline steam
distillation and back titration against standard acid (1).
MeBr oxidation was measured by monitoring its disappearance from the
gas phase of the vials using a Shimadzu GC-14 gas chromatograph. To
account for abiological MeBr hydrolysis and MeBr sorption to vials,
butyl rubber stoppers, and soil, control vials with and without soil
were set up containing 1% (vol/vol) acetylene, a specific
mechanism-based inactivator of AMO (10). The amount of MeBr
in the vials was determined by comparison to standards of known amounts
of MeBr prepared in 74-ml vials containing sterile soil and either 5 ml
(saturated) or 3 ml (unsaturated) of water. Because the amounts of
water in the vials differed between the saturated and unsaturated soil
treatments, slightly different concentrations of MeBr developed in the
aqueous phases. For example, when 10 µmol of MeBr was added to vials,
the aqueous-phase concentrations were calculated to be 0.47 and 0.51 mM
for unsaturated and saturated conditions, respectively. Preliminary
studies established that the presence of soil had no significant impact
on the partitioning of MeBr between the aqueous and gaseous phases and
that 
10% of the MeBr that disappeared during incubations was AMO
independent (i.e., acetylene insensitive). We concluded that some
abiological hydrolysis or sorption of MeBr occurred in our experimental
system, and these values were subtracted from those measured in the
experimental treatments without acetylene. No attempt was made to
distinguish between MeBr disappearance due to sorption and that due to
abiological hydrolysis. In all assays, the amount of MeBr transformed
was expressed as total micromoles inclusive of that amount which
partitioned into the liquid and soil phases.
Effects of intact soil on NO2
production.
The presence of Willamette silt loam soil affected the
rate of NH3 oxidation by N. europaea under both
unsaturated and saturated conditions. When incubations were conducted
at WHC with 10 mM NH4+,
NO2 production occurred at ~30% of the
rate of the minus-soil control and was restored to 
85% of the
minus-soil rate by increasing the concentration of
NH4+ to 40 mM (Fig.
1a). When N. europaea was
incubated with the same weight of soil at approximately two-thirds of
WHC, NO2 production occurred at 15 to 35% of
the minus-soil rate, and a fourfold increase in
NH4+ increased the rate to that of the
minus-soil control (Fig. 1b). Under unsaturated conditions, we noted
occasionally that 40 mM NH4+ would not
completely compensate for the effect of soil on
NO2 production (55 to 60% of the
minus-soil rate). When this phenomenon was observed, larger additions
of NH4+ (50 or 60 mM) did not correct the
problem. To avoid the possibility of either
NH4+ or salt stresses on N. europaea, 40 mM NH4+ was used routinely as
our maximum NH4+ level for unsaturated
conditions. The presence of Willamette silt loam substantially reduced
the concentrations of NH4+ in soil solution
from 40 and 10 mM to approximately 10 and 1 mM, respectively,
regardless of soil water content. Using a Ks value of 4.8 mM for NH4+ oxidation at pH 7.0 (the pH of a 1:1 [vol/wt] soil suspension) and the concentrations of
NH4+ experimentally determined in soil
solutions, theoretical estimates of the rates of
NO2 production matched reasonably well those
experimentally determined, i.e., 24 to 26% and 73 to 80% of the rates
expected in the absence of soil at pH 7.2 and 10 or 40 mM
NH4+, respectively.
|
), 20 mM NH4+
(
), or 40 mM NH4+ (
). The minus-soil
control contained 10 mM NH4+ and the same
amount of cells in either 5 or 3 ml of buffer plus water (
). Error
bars represent the standard deviations of the means of three analytical
replicates. Most standard deviations were <5% of the means and are
not shown.
Effects of soil water content on MeBr oxidation and
NO2 production.
N. europaea
oxidized MeBr under both unsaturated and saturated soil conditions. The
rates of MeBr disappearance were constant for approximately 12 h
and declined to zero between 24 and 36 h (Fig.
2). Because MeBr oxidation was not
sustainable at >0.5 mM aqueous concentration
(MeBraq), subsequent experiments were restricted
to three aqueous concentrations of 0.13, 0.25, and 0.51 mM MeBr
(saturated) or 0.12, 0.24, and 0.47 mM MeBr (unsaturated). As mentioned
earlier, there were slight differences between the aqueous-phase
concentrations in the saturated and unsaturated experiments because
different amounts of water were used. Experiments were conducted to
compare the effect of soil on NO2 production
and MeBr oxidation under water-saturated (Table
1) and water-unsaturated (Table
2) conditions. Saturated soil reduced NO2 production by 10 mM
NH4+-MeBr combinations to 29, 23, and 17%,
respectively, of their corresponding minus-soil values while also
reducing the amounts of 0.24 and 0.47 mM MeBr oxidized to 64 and 43%,
respectively, of their corresponding minus-soil values (Table 1).
Saturated soil reduced NO2 production by 40 mM NH4+-MeBr combinations to a much lesser
degree than it did the 10 mM NH4+-MeBr
combinations, and it increased the amounts of MeBr oxidized by 25, 41, and 33% over the corresponding minus-soil values. The interactions
between saturated soil, NH4+, and MeBr
concentrations were quite striking for both
NO2 production and MeBr oxidation. For
example, 40 mM NH4+ completely compensated for
the negative effect of saturated soil on the amount of MeBr oxidized by
the 0.24 mM MeBr-10 mM NH4+ combination (9.6 versus 8.0 µmol mg [dry weight] of cells1). In
contrast, 40 mM NH4+ only partially compensated
for the effect of saturated soil on the 0.47 mM MeBr-10 mM
NH4+ combination (9.6 versus 12.8 µmol mg
[dry weight] of cells1) despite increasing the rate of
NO2 production by almost the same degree
(fourfold) as it did in the presence of 0.24 mM MeBr.
|
, minus soil and plus 1% acetylene;
|
|
production by 10 mM NH4+-MeBr combinations more severely than
did saturated soil (11 to 12.5% of the corresponding MeBr-minus-soil
combinations) (Table 2). Despite the severe reductions in
NO2 production, oxidations of 0.13 and 0.25 mM MeBr were unaffected, while the amount of 0.51 mM MeBr oxidized was
reduced to 57% of the corresponding minus-soil treatment (9.6 versus
16.8 µmol mg [dry weight] of cells1). In the presence
of 40 mM NH4+, unsaturated soil lowered
NO2 production by the 40 mM
NH4+-MeBr combinations more than occurred in
the saturated-soil experiment (Table 1). The amounts of MeBr oxidized,
however, were increased substantially (60 to 70%) above the minus-soil
values at each of the three MeBr concentrations.
The results of this study clearly reveal the potential of the
NH3 oxidizer N. europaea to oxidize MeBr under
intact soil conditions. Although the rates of oxidation of halogenated
hydrocarbons by soilborne populations of NH3 oxidizers are
not currently available in the literature, we intentionally used
relatively low-density cell suspensions of N. europaea in an
attempt to generate rates of MeBr oxidation and
NO2 production that could be placed into
context with rates documented elsewhere in the microbial ecology
literature. For example, we measured rates of soilborne
NO2 production of 25 to 60 nmol g of
soil1 h1, which are on the high end of the
range of values generally accepted for nitrification potentials of
actively nitrifying soils (1 to 100 nmol of N g of soil1
h1) (16). The rates of MeBr oxidation fell
into the range of ~5 to 10 nmol of MeBr transformed g of
soil1 h1. These values are similar to rates
of MeBr degradation reported for a methanotrophic peat exposed to MeBr
concentrations similar to those used in this study (17).
They are much greater than the rates of 1 to 3 nmol of MeBr transformed
g of soil1 day1 reported for a fumigated
agricultural soil (15) and of ~2 pmol g of
soil1 h1 for degradation of atmospheric
levels of MeBr by a forest soil (7).
Our data highlight how soil physical and chemical properties can modify
the characteristics of MeBr oxidation by NH3 oxidizers through their influence on the bioavailability of the two substrates. Previous studies illustrated that exchangeable soil acidity was the
primary factor influencing NH3 availability to N. europaea under soil slurry conditions (8). In the
present study, which was conducted with acid-neutralized soil, it
became quite clear that much larger additions of
NH4+ were needed to compensate for the effect
of structurally intact soil on NO2 production
than was apparent under slurry conditions. Our data clearly indicate
the interactive effect of soil water content on cooxidation of a
halogenated hydrocarbon through its differential influence on the
availability of NH4+ and MeBr (i.e., two
substrates with quite different chemical properties). In general, there
was a greater negative impact of unsaturated soil on
NO2 production than on MeBr oxidation, while
saturated soil had a greater inhibitory effect on MeBr oxidation than
on NO2 production. These findings are
consistent with the observation that nitrification declines as soil
water content is lowered because of increased restrictions on the
diffusion of NH4+ to the sites of the
NH3-oxidizing bacteria (23). Furthermore, it is
not difficult to conceptualize that MeBr oxidation might be restricted
by diffusion at higher soil water content, provided the cooxidative
process is not already NH4+ limited. Indeed,
several closely related studies have shown that CH4
consumption by soil occurs optimally at 20 to 40% of WHC and usually
declines as water content is raised beyond this range (4, 27,
28). In one of these soil studies, the optimum water content for
CO2 production was found to be significantly higher than
for CH4 consumption (4). The authors concluded that because CH4 consumption was reliant on gaseous
diffusion, it would be negatively affected by an increase in soil water
content, while soil respiration would respond positively because of its reliance on the diffusion of water-soluble substrates. More studies are
required to establish to what extent differences among soils in their
properties of NH4+ generation, adsorption, and
availability might interact with water-holding characteristics to
influence the relationships between NH3 and gaseous
hydrocarbon oxidation.
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ACKNOWLEDGMENTS |
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This work was supported by the Oregon Agricultural Experiment Station and by EPA grant R821405 to P.J.B. and D.J.A. Additional support was provided to K.N.D. through the Department of Microbiology of Oregon State University and the N. L. Tarter Fellowship.
We express appreciation to David Myrold and Chris Yeager for constructive comments on early drafts of the manuscript.
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FOOTNOTES |
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* Corresponding author. Mailing address: Department of Microbiology, 220 Nash Hall, Oregon State University, Corvallis, OR 97331. Phone: (541) 737-1844. Fax: (541) 737-0496. E-mail: bottomlp{at}ucs.orst.edu.
Technical paper no. 11,573 of the Oregon Agricultural Experiment Station.
Present address: Department of Biological Sciences, University of
Alaska Anchorage, Anchorage, AK 99508.
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Applied and Environmental Microbiology, June 2000, p. 2636-2640, Vol. 66, No. 6
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Copyright © 2000, American Society for Microbiology. All rights reserved.
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