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Applied and Environmental Microbiology, May 1999, p. 2246-2249, Vol. 65, No. 5
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Removal of Dibenzofuran,
Dibenzo-p-Dioxin, and
2-Chlorodibenzo-p-Dioxin from Soils Inoculated with
Sphingomonas sp. Strain RW1
Rolf U.
Halden,
Barbara G.
Halden, and
Daryl F.
Dwyer*
Department of Civil Engineering, University
of Minnesota, Minneapolis, Minnesota 55455
Received 19 January 1999/Accepted 23 February 1999
 |
ABSTRACT |
Removal of dibenzofuran, dibenzo-p-dioxin, and
2-chlorodibenzo-p-dioxin (2-CDD) (10 ppm each) from soil
microcosms to final concentrations in the parts-per-billion range was
affected by the addition of Sphingomonas sp. strain RW1.
Rates and extents of removal were influenced by the density of RW1
organisms. For 2-CDD, the rate of removal was dependent on the content
of soil organic matter (SOM), with half-life values ranging from
5.8 h (0% SOM) to 26.3 h (5.5% SOM).
| |
TEXT |
Diaryl ether compounds include
several chemicals (e.g.,
2,3,7,8-tetrachlorodibenzo-p-dioxin) that are recognized as
toxic pollutants and that persist in soils, in part, by sorbing onto organic matter (7, 20) and, to a lesser extent, onto mineral surfaces (12). Aerobic bacteria that degrade
dibenzo-p-dioxin (DD), dibenzofuran (DF), carboxydiphenyl
ether, and some halogenated derivatives of these chemicals have been
isolated (9, 11, 16, 17, 26, 32, 33). Researchers have
primarily focused their studies on the activities of these bacteria in
liquid culture (10, 18, 19). In one case, degradation of DD
and DF in soil microcosms to which Sphingomonas sp. strain
RW1 was introduced was observed (25). To determine whether
these and similar bacteria have potential for use in in situ
bioremediation, data concerning the fate and activity of the bacteria
in soils are needed in conjunction with data on the bioavailability of
target chemicals (14). These factors were addressed in this
study by using soil microcosms and Sphingomonas sp. strain
RW1, a bacterium that mineralizes DD and DF as growth substrates and
transforms certain mono- and dichlorinated analogs of these chemicals
as cometabolic reactions (32).
Bacterial growth conditions.
Strain RW1 (no. 6014; German
Collection of Microorganisms and Cell Cultures, Braunschweig, Germany)
was tagged with a Tn5 suicide donor system, pNMBH20, that
encodes genes for resistance to kanamycin and for
catechol-2,3-dioxygenase (24). This enzyme converts catechol
to muconic acid semialdehyde, a product having a bright yellow color,
and does so at a higher rate than the catechol-2,3-dioxygenase present
in strain RW1. Strain RW1 was routinely grown in the dark (200 rpm,
30°C) by using M9 minimal medium (23) supplemented with
trace elements (31) and DF (1 g/liter) as the sole
substrate. Solid medium contained agar (15 g/liter) with benzoate (5 mM) as the sole substrate plus kanamycin (50 µg/ml).
Soils.
A pale brown, sandy Zimmerman soil (B21 horizon, 18 to
38 cm below surface) was obtained from a previously cultivated field in
the Cedar Creek Natural History Area, Minn. A dark soil (A horizon) was
obtained from Fort Snelling State Park, St. Paul, Minn. Soils were
sieved (2-mm mesh) and stored at 4°C. A quartz sand (Jordan) was
obtained from the Minnesota Frac Sand Co., Jordan, Minn. (Table
1). The sand was sieved to obtain a
fraction with a particle size distribution of 0.15 to 0.6 mm. Cedar
Creek soil was blended with Jordan sand and Fort Snelling soil to
obtain various contents of organic matter, ranging from 0 to 5.5%
total organic carbon (TOC). Mixtures were air dried, and water was
added to a level equal to 60% field capacity.
Soil microcosm studies.
Microcosms consisted of either serum
bottles (100 ml) plus soil (20 g [dry weight] of soil) or vials (4 ml) plus soil (2 g [dry weight] of soil). DD,
2-chlorodibenzo-p-dioxin (2-CDD) (99% purity; Chemservice,
West Chester, Pa.), and DF (Sigma, St. Louis, Mo.) were added to
individual microcosms from methanolic stocks 24 h prior to
inoculation with strain RW1; strain RW1 does not use methanol as
substrate. Bacteria were grown to late log phase, collected by
centrifugation (5,000 × g, 20 min), washed twice in
saline (0.85% NaCl), and added to the microcosms during mixing. The
microcosms were covered with Parafilm and incubated (21°C) in the
dark. Densities of strain RW1 and concentrations of DD, DF, and 2-CDD
were determined periodically by sacrificing individual microcosms. Data
are reported as averages of triplicate determinations.
Bacteria were extracted from soil in an extraction solution (1 ml/g
[dry weight] of soil [
21]) by agitation on a wrist
action
shaker (30 min). Extracts were serially diluted in saline;
aliquots
were then spread on solid medium and incubated (30°C, 7 days).
Resultant colonies were sprayed with a solution of catechol (100
mM) to identify strain RW1. The detection limit was 10
4
CFU/g (dry weight) of soil; the efficiency of recovery was 87%.
DD, DF, and 2-CDD were extracted from soil by addition of acetonitrile
(2% H
3PO
4, 1 ml/g [dry weight] of soil) and
agitation
(60 min). Particles were settled out (10 min), and the
supernatant
was passed through a PTFE membrane filter (0.2 µm).
Aliquots were
analyzed by high-pressure liquid chromatography with a
Waters
LC Module I equipped with a reversed-phase Waters Nova-Pak
phenyl
column (3.9 by 150 mm; particle size, 4 µm) and a Waters 996 photodiode
array detector (
191-320 nm). An isocratic
solvent of acidified,
distilled water (0.1%
H
3PO
4) and acetonitrile (50:50) was used.
Injection was by automatic sampler; volumes ranged from 10 to
200 µl.
Peaks and concentrations were identified by comparison
to known
standards. Recoveries of DF, DD, and 2-CDD ranged from
89 to 100%;
detection limits were 50 ppb. As appropriate, concentrations
of
chemicals were fit to either a pseudo-first-order rate model,
dP/
dt =
k
P, or a second-order rate
model,
dP/
dt =
k2BP,
where
P
is the concentration of substrate,
B is the concentration
of
biomass,
t is time,
k is the
pseudo-first-order rate constant,
and
k2 is the
second-order rate
constant.
Survival of RW1.
Densities of strain RW1 (4 × 108 CFU/g [dry weight] of soil) decreased exponentially
in soils without substrate amendment at a rate (0.156 ± 0.007 day
1) corresponding to a half-life of 4.4 days. In the
presence of DF (10 ppm), the decrease in density was less rapid, at a
rate (0.093 ± 0.034 day1) corresponding to a
half-life of 7.5 days. The same rate was observed for several initial
densities of the bacterium (4 × 105, 4 × 106, and 4 × 107 CFU/g [dry weight] of
soil). In contrast, the presence of DD (10 ppm) did not affect the
survival of strain RW1; the density of the bacterium decreased at a
rate of 0.148 ± 0.004 day1, which is similar to
that observed in soils without substrate amendment. Densities of strain
RW1 decreased dramatically in soils containing 2-CDD (10 ppm), at a
rate (0.782 ± 0.045 day1) corresponding to a
half-life of 0.9 days (data not shown).
Degradation of diaryl ether compounds.
DF was removed from
soils containing strain RW1 (Fig. 1a).
The extent of removal was dependent on the initial density of the bacteria: a density of 4 × 107 CFU/g (dry weight) of
soil resulted in complete removal of DF (<50 ppb) in 7 days. At lower
densities, removal of DF was incomplete after 28 days of incubation. DD
was also removed by strain RW1 (Fig. 1b); however, relatively higher
densities of cells were required. At the highest density tested
(109 CFU/g [dry weight] of soil), 90% of DD was degraded
within 24 h; after 3 days, the concentration of DD was below the
detection limit (50 ppb). In comparison to DD, 2-CDD was somewhat more
persistent in soil (Fig. 1c).
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FIG. 1.
Removal of DF (a) from soils inoculated with RW1 at
densities of 4 × 105 ( ), 4 × 106
( ), and 4 × 107 ( ) CFU/g (dry weight) of soil
and of DD (b) and 2-CDD (c) from soils inoculated with RW1 at densities
of 107 ( ), 108 ( ), and 109
( ) CFU/g (dry weight) of soil. Control microcosms ( ) received
substrates but not RW1.
|
|
Removal of 2-CDD by strain RW1 was tested further with soils having
varying levels of organic matter; concentrations of 2-CDD
were
monitored over time (data not shown) and used to calculate
rates of
degradation (Table
2) with the
pseudo-first-order rate
model. Rates decreased when comparatively more
soil organic matter
(SOM) was present; half-life values ranged from 5.8 to 26.3 h.
The influence of organic matter became more apparent
when rates
of degradation for 2-CDD were plotted against SOM (Fig.
2a) and
TOC (Fig.
2b). Strong
correlations that clearly demonstrated the
negative impact of
increasing amounts of organic matter on removal
rates for 2-CDD were
observed.
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TABLE 2.
Pseudo-first-order rate coefficients and half-life values
for 2-CDD in soils containing RW1 (109 CFU/g [dry
weight] of soil) and various amounts of SOM
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FIG. 2.
Correlations between pseudo-first-order rates of
degradation of 2-CDD by RW1 with SOM (a) and TOC (b). Error bars
indicate standard errors obtained by curve fitting.
|
|
Data in Fig.
2b were used to obtain densities of strain RW1 that would
be required to degrade 10, 50, 90, and 99% of 2-CDD
in soils
containing various concentrations of organic matter (Fig.
3). Removal rates for 2-CDD were
estimated with the correlation
in Fig.
2b and the estimated rates
divided by cell density (10
9 CFU/g [dry weight] of soil)
to obtain specific transformation
rates (
k2) per
cell. The second-order rate model was then used
to calculate cell
densities for each percentage of TOC. This exercise
indicated that as
the TOC content of soil increased, relatively
more biomass was required
to achieve the same extent of removal
of 2-CDD. For example, it was
necessary to add approximately three
times more biomass to soil
containing 4% TOC than to soil containing
0% TOC to effect removal of
90% 2-CDD.
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FIG. 3.
Calculated densities of RW1 (CFU/g [dry weight] of
soil [dws]) that would be required to achieve 2-CDD removal of 10, 50, 90, and 99% within 24 h in soils with TOC ranging from 0 to
4%.
|
|
This study demonstrates three points with respect to the potential use
of diaryl ether-degrading bacteria for in situ bioremediation.
(i)
Bacteria with catabolic pathways for diaryl ethers, as represented
by
strain RW1, can survive and degrade diaryl ether compounds
in soil.
(ii) Diaryl ethers can exist as bioavailable chemicals
in soils, at
least under the conditions used in this study. (iii)
Soils having
similar densities of diaryl ether-degrading microorganisms,
but
relatively higher levels of TOC, may exhibit relatively lower
rates of
degradation of these diaryl ether
compounds.
It was interesting that the three diaryl ether compounds used in this
study exerted different influences on the survival of
strain RW1.
Although degradation of DF prolonged the persistence
of strain RW1, an
increase in bacterial density was not observed,
possibly due to a lack
of essential nutrients. This suggestion
was supported by the
observation that nutrient supplementation
increased the density of a
second diaryl ether-degrading bacterium
that was added to Cedar Creek
soil (
15). In contrast, degradation
of DD did not prolong
the persistence of strain RW1. This may
be related to a difference in
utilization rates for DF and DD
by strain RW1, as was noted in a
similar study (
25). In pure
culture, the turnover rate for
DD (280 µmol h
1 g of protein
1) is
relatively slower than the turnover rate for DF (562 µmol
h
1 g of protein
1) (data from reference
32) and may not be sufficient to prolong
survival of
the
microorganism.
The survival of strain RW1 was adversely affected by the presence of
2-CDD, suggesting that 2-CCD and/or its metabolites were
toxic to the
bacterium. Dioxins are typically neither toxic nor
inhibitory to
microorganisms and have no observable effect on
soil respiration
(
5), microbial activity, and diversity (
2).
However, during transformation of 2-CDD, 4-chlorocatechol is observed
as a dead-end product (
32). Chlorinated phenolics are used
as
bactericides (
27), and chlorocatechols are inhibitors of
key
catabolic enzymes (
4,
29,
33). Thus, production of
4-chlorocatechol
may have accounted for the relatively poor survival of
strain
RW1. It appears that by producing toxic intermediates, diaryl
ether-degrading microorganisms could be negatively impacted during
attempts to elicit in situ bioremediation. Judicious engineering
of
microorganisms for complete mineralization of diaryl ethers,
as has
been suggested previously (
14,
32), may offer a way
around
this
dilemma.
It is apparent that SOM controlled the rate at which 2-CDD was removed
from soil by strain RW1. Bioavailability of hydrophobic
pollutants can
potentially limit degradation of organic chemicals
in soils (
1,
3,
30) and is typically controlled by physicochemical
processes
including diffusion, sorption/desorption, and dissolution
(
6,
22,
28). Relatively nonpolar chemicals such as dioxins
primarily
partition into SOM rather than mineral surfaces (
7,
8,
13,
20). The correlation between increasing TOC and
decreasing rates
of removal of 2-CDD suggests that the rate of
degradation was limited
by sorption/desorption processes. Thus,
as TOC content increases, the
lower bioavailability of 2-CDD requires
that the amount of biomass be
increased to elicit effective degradation
(Fig.
3). For soils with
elevated levels of TOC, this means that
the relative cost of
remediation via bioaugmentation may be higher,
due to the necessity of
adding relatively more
microorganisms.
| |
ACKNOWLEDGMENTS |
We thank R. Wittich for RW1, N. McClure for pNMBH20, and the
Research Analytical Laboratory, University of Minnesota, for soil analyses.
This research was supported by the National Science Foundation
(BCS-9318788). Financial support for R.U.H. was provided in part by a
University of Minnesota Doctoral Dissertation Fellowship.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Civil Engineering, 122 Civil Engineering Building, University of
Minnesota, 500 Pillsbury Dr., S.E., Minneapolis, MN 55455. Phone: (612)
625-8582. Fax: (612) 626-7750. E-mail:
dwyer003{at}tc.umn.edu.
Present address: Lawrence Livermore National Laboratory,
Environmental Restoration Division, Livermore, CA 94550.
| |
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Applied and Environmental Microbiology, May 1999, p. 2246-2249, Vol. 65, No. 5
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
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Halden, R. U., Tepp, S. M., Halden, B. G., Dwyer, D. F.
(1999). Degradation of 3-Phenoxybenzoic Acid in Soil by Pseudomonas pseudoalcaligenes POB310(pPOB) and Two Modified Pseudomonas Strains. Appl. Environ. Microbiol.
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