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Applied and Environmental Microbiology, December 2000, p. 5527-5532, Vol. 66, No. 12
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Utilization of Dimethyl Sulfide as a Sulfur Source
with the Aid of Light by Marinobacterium sp. Strain
DMS-S1
Hiroyuki
Fuse,1,*
Osamu
Takimura,1
Katsuji
Murakami,1
Yukiho
Yamaoka,1 and
Toshio
Omori2
Chugoku National Industrial Research
Institute, 2-2-2 Hirosuehiro, Kure, Hiroshima
737-0197,1 and Biotechnology Research
Center, University of Tokyo, Bunkyo-ku, Tokyo
113-9657,2 Japan
Received 5 June 2000/Accepted 2 October 2000
 |
ABSTRACT |
Strain DMS-S1 isolated from seawater was able to utilize dimethyl
sulfide (DMS) as a sulfur source only in the presence of light in a
sulfur-lacking medium. Phylogenetic analysis based on 16S ribosomal DNA
genes indicated that the strain was closely related to
Marinobacterium georgiense. The strain produced dimethyl sulfoxide (DMSO), which was a main metabolite, and small amounts of
formate and formaldehyde when grown on DMS as the sole sulfur source.
The cells of the strain grown with succinate as a carbon source were
able to use methyl mercaptan or methanesulfonate besides DMS but not
DMSO or dimethyl sulfone as a sole sulfur source. DMS was transformed
to DMSO primarily at wavelengths between 380 and 480 nm by heat-stable
photosensitizers released by the strain. DMS was also degraded to
formaldehyde in the presence of light by unidentified heat-stable
factors released by the strain, and it appeared that strain DMS-S1 used
the degradation products, which should be sulfite, sulfate, or
methanesulfonate, as sulfur sources.
| |
TEXT |
Dimethyl sulfide (DMS) released from
the sea is an important compound in global sulfur circulation
(28) and global climate regulation (4). DMS is
generated by the degradation of dimethylsulfoniopropionate, which is
present in many species of marine algae and plants, including dinoflagellates and coccolithophores (20). However, more
than 10 times the amount of DMS released to the atmosphere is degraded in the sea by microorganisms (23). DMS is degraded or
transformed by terrestrial and marine microorganisms via methyl
mercaptan (MM) or dimethyl sulfoxide (DMSO). Some strains of sulfur
oxidizers (5, 19, 29, 32, 33) and methylotrophs (7, 9, 30, 38) degrade DMS via MM. Ammonia oxidizers (18),
methanotrophs (11), algae (12), and some strains
of phototrophs (15, 31, 36) transform DMS to DMSO. Some
terrestrial heterotrophic bacterial strains also have been found to
degrade or transform DMS via DMSO. Comamonas acidovorans
DMR-11 (originally Pseudomonas acidovorans) isolated from
peat biofilters transformed DMS to DMSO in the medium containing other
organic carbon sources, such as sodium malate (37). A
dibenzothiophene-desulfurizing bacterium, Rhodococcus sp.
strain SY1, was able to degrade DMS in the oxidative pathway via DMSO,
dimethyl sulfone, methanesulfonate, and sulfate, and genes encoding an
enzyme that oxidized DMS to DMSO have been cloned from
Acinetobacter sp. strain 20B, which was able to grow on DMS as the sole sulfur source (17, 26). Recently, several marine isolates in the 
subclass of the class Proteobacteria
have been found to be able to transform DMS to DMSO or MM
(14). There have been no reports on marine heterotrophic
isolates other than the subclass of the class
Proteobacteria that are able to degrade DMS aerobically.
This paper reports the isolation and characterization of a marine
heterotrophic bacterium that belongs to the 
subclass of the class
Proteobacteria and is able to utilize DMS as the sole sulfur
source only in the presence of light.
Isolation and characterization of strain DMS-S1.
The basal
medium (NSYE) for isolation and cultivation of strain DMS-S1 contained
25 g of NaCl, 0.7 g of KCl, 50 mg of
KH2PO4, 1 g of
NH4NO3, 0.2 g of MgCl2
· 6H2O, 20 mg of CaCl2 · 2H2O, 5 mg of FeEDTA, 1 g of Tris, 5 mg of yeast
extract, and 5 g of sodium succinate in 1 liter of distilled
water. The final pH was adjusted to 7.7 to 8.0 with NaOH solution. The
basal medium was autoclaved at 110°C for 10 min. ZoBell medium
(27) with 1.5% agar was also used for isolation by
streaking. Isolation and cultivation were done at 21°C with shaking
at 100 rpm under illumination (45 to 57 µmol · m
2 · s1) provided by 20-W
fluorescent lamps. Strain DMS-S1 was isolated from a marine sample
taken from Edauchi Bay (Hiroshima, Japan) in November 1992. The cells
of strain DMS-S1 are gram-negative, non-spore-forming, aerobic, short
rods with single polar flagella. They are oxidase positive, catalase
positive, and O-F test negative, and they require a seawater base for
growth. The activities of DNA hydrolysis and gelatin hydrolysis were
not found. The quinone type of the cells was Q-8. The G+C content of
the DNA was 56 mol%. To test growth on various carbon sources, carbon
sources were added to SWNC medium (1 g of
NH4NO3, 0.5 g of
KH2PO4, 5 mg of FeEDTA, and 10 mg of yeast
extract in 1 liter of filtered seawater, pH 7.7 to 8.0) or to NSYE
medium without sodium succinate and containing 2 to 10 mM
Na2SO4. The strain was cultured in 25- by 200-mm (71-ml capacity) test tubes containing 20 ml of media with Teflon-lined screw caps. Strain DMS-S1 was able to utilize succinate, acetate, ethanol, propanol, and butanol as carbon sources. The strain
was not able to utilize glucose, glycerol, methanol, DMS, DMSO,
dimethyl sulfone, methanesulfonate, diethyl sulfide,
tetrahydrothiophene, diethyl sulfone, ethanesulfonate, methionine, or
(2-carboxyethyl)dimethylsulfonium chloride as carbon sources.
A 16S ribosomal DNA sequence containing 1,478 bp of strain DMS-S1 was
analyzed as described previously (11). Strain DMS-S1 was
closely related to Marinobacterium georgiense, which was
isolated from a marine pulp mill effluent enrichment culture
(13). The similarity of the 1,423 bp of strain DMS-S1 to the
M. georgiense sequence was 98.2%. The characteristics of
strain DMS-S1 observed here resembled those of M. georgiense
reported by Gonzalez et al. (13) except for the utilization
of glucose, glycerol, and methanol as carbon sources, though the media
used for the tests were different. M. georgiense ATCC 700074 was not able to grow on DMS as a sulfur source under the same culture
conditions as strain DMS-S1 within 5 days after inoculation. Based on
these findings, strain DMS-S1 was identified as a
Marinobacterium sp.
Growth of strain DMS-S1 on organic sulfur compounds as sole sulfur
sources.
The growth of strain DMS-S1 on DMS as a sulfur source was
compared with growth on Na2SO4. For the
experiments assessing growth on organic sulfur compounds as sulfur
sources, a sulfur source was added to the medium after filter
sterilization with a 0.2-µm-pore-size membrane filter. Precultured
strain DMS-S1 in sulfate-containing medium was inoculated after
dilution with the basal medium. Growth of the strain was monitored at
600 nm with a Spectronic 20D spectrophotometer (Milton Roy Co.,
Rochester, N.Y.). Cultures were always run in pairs under the same
conditions, and values for growth are shown here as the means for the
two cultures. We have not analyzed the sulfur concentration of basal
medium exactly. However, the results of the growth experiment (Fig.
1) showed that the sulfur compounds present in the basal medium and carried over from preculture were less
effective than 0.2 µM Na2SO4. The maximum
growth of the strain on 4 µmol of DMS, which is calculated to be 0.16 mM in the medium after equilibrium (6), was less than that
on 400 nmol of Na2SO4 (20 µM in the medium)
and more than that on 40 nmol of Na2SO4 (2 µM
in the medium) (Fig. 1). This suggests that the strain requires over
10-fold more DMS than Na2SO4 to support growth.
Effects of light on the growth of the strain were examined (Fig.
2). The strain needed light only when DMS
was the sole sulfur source.
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FIG. 1.
Growth of strain DMS-S1 utilizing
Na2SO4 or DMS as a sulfur source. Four
nanomoles of Na2SO4 ( ), 40 nmol of
Na2SO4 ( ), 400 nmol of
Na2SO4 ( ), 400 nmol of DMS ( ), 4 µmol
of DMS ( ), or 40 µmol of DMS ( ) was added to 71-ml test tubes
containing 20 ml of NSYE medium. The concentrations in the medium after
equilibrium were calculated to be as follows: 0.2 µM
Na2SO4 (), 2 µM
Na2SO4 (), 20 µM
Na2SO4 (), 16 µM DMS (), 160 µM DMS
(), and 1.6 mM DMS (). ×, NSYE medium without added sulfur
sources. OD600, optical density at 600 nm.
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FIG. 2.
Growth of strain DMS-S1 with or without light. Twenty
milliliters of medium was used. ×, basal medium without added sulfur
sources; , basal medium with 2 mM Na2SO4;
, basal medium with 16 mM DMS (total, 400 µmol in a tube); ,
ZoBell medium.
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Organic sulfur compounds utilized as sulfur sources by the strain are
shown in Table
1. When MM gas was added
to the test
tubes, screw caps with a valve for a syringe were used.
This strain
was able to utilize diethyl sulfide but not
di-
n-propyl sulfide
or di-
n-butyl sulfide. It was
not able to grow on DMSO or dimethyl
sulfone but was able to grow on MM
as a sulfur source, unlike
Rhodococcus sp. strain SY1 and
Acinetobacter sp. strain 20B, which
were able to grow on
DMSO and dimethyl sulfone but not on MM as
a sulfur source (
17,
26). Strain DMS-S1 also utilized alkanesulfonates
as a sulfur
source. MM and methanesulfonate were utilized even
in the absence of
light.
Metabolites from DMS produced by growth of strain DMS-S1.
The
culture supernatant of strain DMS-S1 grown on DMS was analyzed by gas
chromatography-mass spectrometry (GC-MS). For GC-MS, a JMS Automass 150 (JEOL, Tokyo, Japan) or a QP-5000 (Shimadzu, Kyoto, Japan) with a 30-m
capillary column (DB-5; J&W Scientific, Folsom, Calif.) was used. The
obtained fragmentation of a metabolite from DMS occurring at
m/z (%) 78 (58) and 63 (100) had a pattern similar to that
from authentic DMSO, whose fragments were at m/z (%) 78 (63) and 63 (100). The metabolite was identified as DMSO. Residual DMS
and accumulated DMSO produced by the growth of strain DMS-S1 on DMS
were quantified by GC with a flame photometric detector as described
previously (12). DMSO was the main product when strain
DMS-S1 was grown on DMS (Fig. 3).
Accumulation of DMSO in the medium without inoculation of the strain
was negligible. There were no metabolites corresponding to dimethyl
sulfone on the gas chromatogram.
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FIG. 3.
Growth and DMSO transformed from DMS by strain DMS-S1.
The concentration of DMS in the medium was calculated to be 1.7 mM at
the start. Growth of strain DMS-S1 is expressed as the optical density
at 600 nm (OD600) (). DMSO () and residual DMS ()
are expressed in micromoles. Values are means for two samples.
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The factor for transforming DMS to DMSO in the culture was
investigated. Strain DMS-S1 was cultured in NSYE medium containing
20 µM Na
2SO
4 for several days. The broth was
separated into supernatant
and cells by centrifugation (6,300 to 9,800 ×
g , 10 min) and
filtration with 0.2-µm-pore-size
Nucleopore filters. The supernatant
was subjected to ultrafiltration
using USY-1 (Advantec, Tokyo,
Japan), which separated molecules larger
than 10,000. The retained
concentrate of the ultrafiltration was made
up with NS buffer
(25 g of NaCl, 0.7 g of KCl, 0.2 g of
MgCl
2 · 6H
2O, 20 mg of
CaCl
2 · 2H
2O, and 0.5 g of Tris in
1 liter of distilled water, pH 7.7)
to the volume of the supernatant
subjected to ultrafiltration.
Cells separated by centrifugation were
washed with NS buffer and
suspended in the buffer at the same volume as
the broth. The factor
was found in the culture supernatant but not in
the cells and
was stable to heat treatment for 5 min at 105°C. The
molecular
weight was lower than 10,000 (Fig.
4). An absorption spectrum
of the culture
supernatant which was prepared from the culture
grown on 40 µM
Na
2SO
4 was measured in a Hitachi 150-20 spectrophotometer.
The culture supernatant of the strain was almost
colorless, though
its absorption spectrum had two local maxima at about
340 and
412 nm (Fig.
5). Wavelengths of
fluorescent lamps needed for the
oxidation of DMS to DMSO in the heated
culture supernatants were
investigated by using seven optical filters
purchased from Kenko.
Absorption characteristics of the filters are
shown in Fig.
6.
The oxidation of DMS
occurred predominantly between 380 and 480
nm (Fig.
7). This wavelength is almost coincident
with the result
for seawater (
21). These facts suggested
that the factor for
transforming DMS to DMSO was a photosensitizer
found in algae
and seawater (
2,
12).
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FIG. 4.
Transformation of DMS by culture components of strain
DMS-S1. The strain was grown in NSYE medium with 20 µM
Na2SO4, and the culture medium was separated
into components as described in the text. Forty-five micromoles of DMS
was added to 4.5 ml of culture component solution in 22-ml vials (7.8 mM DMS in the solution). (A) Whole culture medium containing cells of
strain DMS-S1 (), culture supernatant ( ), and cells in buffer
(). (B) Whole culture medium containing cells heated for 5 min at
105°C () and culture supernatant heated for 5 min at 105°C
( ). (C) Components of culture supernatant whose molecular weight was
higher than 10,000 in buffer () and culture supernatant after
exclusion of the components with a molecular weight higher than 10,000 ().
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FIG. 7.
DMSO accumulation by heated culture supernatants under
light passed through optical filters. Twenty-nine micromoles of DMS was
added to 3.6 ml of heated (105°C, 5 min) culture supernatants in
22-ml vials. The vials were kept under light passed through optical
filters, and the DMSO that accumulated in the supernatants was
quantified after 4 days. NSYE medium containing 40 µM
Na2SO4 and 29 µmol of DMS was used as a
control and kept under light without a filter. Error bars indicate
standard deviations for three samples.
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Production of formate by the strain was investigated via high-pressure
liquid chromatography after derivatization of products
with
2-nitrophenylhydrazine (
1). When formate in the cultured
medium was to be quantified, 1 to 2 ml of ethanol, butanol, or
propanol
was substituted for 5 g of sodium succinate in 1 liter
of NSYE
medium because large amounts of succinate interfered with
the
derivatization of formate and made the detection of formate
difficult.
Accumulation of formate was detected only in the cultures
grown on DMS
as a sulfur source, and it was negligible in the
cultures grown on
sulfate no matter what alcohol was used as a
carbon source. This fact
suggested that formate was produced during
DMS utilization by this
strain or that DMS might have inhibited
the metabolism of formate
derived from other sources. Formate
accumulated from DMS was about 3 to
5% (mol/mol) of the added
40 µmol of DMS after cultivation for 13 to
18 days. Accumulation
of formate from DMS by the culture grown on
sulfate as a sulfur
source was suppressed in the absence of light (data
not
shown).
In addition, we examined the production of formaldehyde by the culture
components of the strain. A product from DMS was identified
via GC-MS
and quantified via high-pressure liquid chromatography
after
derivatization to formaldehyde-2,4-dinitrophenylhydrazone
(
22). The obtained fragmentation of the derivatized
metabolite
from DMS occurring at
m/z (%) 210 (50), 180 (9),
152 (11), 122
(22), 91 (18), 79 (77), 63 (100), and 51 (91) had a
pattern similar
to that obtained from the authentic
formaldehyde-2,4-dinitrophenylhydrazone,
with fragments at
m/z (%) 210 (47), 180 (11), 152 (8), 122 (20),
91 (22), 79 (93), 63 (100), and 51 (94). Thus, the metabolite
was identified as
formaldehyde. A larger amount of formaldehyde
was accumulated by the
culture supernatant than by the cells in
buffer, and it was also
accumulated by the culture supernatant
heated for 5 min at 105°C,
though formate was not accumulated
by the culture supernatant (Table
2). These facts confirmed that
formaldehyde and formate were produced during DMS utilization
by this
strain. Accumulation of MM was not observed with the production
of
formaldehyde by the heat-treated culture components.
Proposed pathway of DMS degradation by strain DMS-S1.
DMS is
readily oxidized to DMSO by photochemical reactions in the presence of
photosensitizers such as humic acid, methylene blue, and rose bengal
(2), though DMS does not undergo appreciable photo-oxygenation in the absence of photosensitizers. Our results for
DMSO production from DMS suggested that strain DMS-S1 excreted substances that served as photosensitizers. However, formaldehyde, which was an unexpected product in the photosensitizing reaction, was
also produced from DMS by the supernatant of this strain in the
presence of light. DMS is also photooxidized to formaldehyde, sulfur
dioxide, sulfate, and methanesulfonate under light in the presence of
NOX in the air (16, 35). Hatakeyame et al.
(16) reported that methanesulfonate was the main product and
the yield was more than 50%. On the other hand, according to Yin et
al. (35), sulfur dioxide was the main product and the yield
was 62 to 71%. We have not been able to detect sulfite, sulfate, or methanesulfonate in this reaction because high concentrations of salts
prevented detection of these compounds by the methods available to us.
On the other hand, strain DMS-S1 could utilize sulfate and
methanesulfonate as a sulfur source but not DMSO, and production of MM
was not observed in this photooxidation of DMS to formaldehyde.
Therefore, sulfite, sulfate, and methanesulfonate are the most likely
intermediates that serve as sources of sulfur during the utilization of
DMS by this strain. Based on the results and the speculations described
above, we propose the degradation pathway of DMS by
Marinobacterium sp. strain DMS-S1 shown in Fig. 8. This pathway corresponds with the
results of growth experiments. DMS is transformed mainly to unusable
DMSO and only a small percentage of DMS is used for growth, which was
why more than 10 times more DMS than sulfate was required for growth.
The compounds that aid in photolysis of DMS need to be purified to
clarify the photooxidation of DMS by this strain.
Sulfate is assimilated after reduction to sulfite (
25).
Methanesulfonate is decomposed and sulfite is released from it by
monooxygenases in
Methylosulfonomons methylovora
(
8) and
E. coli (
10). Some marine
bacteria belonging to the
subclass
of the class
Proteobacteria are able to release MM from
dimethylsulfoniopropionate
and DMS and incorporate it directly into
methionine (
14,
24).
They use MM in preference to sulfate,
which is present at 10
6- to 10
7-fold-higher
concentrations. The merit of usage of MM is to save
the reducing power
needed for conversion of sulfate to sulfide.
As for strain DMS-S1, the
assimilation of sulfite or methanesulfonate
instead of sulfate could
also provide an energetic advantage,
though they are less effective
than
MM.
The oxidation of DMS to other compounds in the sea plays an important
role in sulfur circulation because the oxidation reduces
the release of
DMS into the air. The reaction decomposing DMS
to formaldehyde is
irreversible, while DMSO oxidized from DMS
can be reduced back to DMS
again (
34). Kieber et al. pointed
out that only 14% of DMS
photolyzed was converted to DMSO and
that the relatively low conversion
was not due to losses of DMSO
(
21). The reaction decomposing
DMS to formaldehyde may play
some role in the loss of DMS. Photolysis
of DMS accounted for
7 to 40% of the total turnover of DMS in the
photic zone of the
equatorial Pacific Ocean (
21). On the
other hand, 88% of the
DMS was photolyzed in the top 10 m of the
water column of the
northern Adriatic Sea (
3). Not only the
quantity but also the
quality of dissolved organic carbon affects the
photolysis of
DMS, as mentioned by Brugger et al. (
3). This
study showed
that marine bacteria excrete substances which could
transform
DMS to DMSO and decompose DMS, releasing formaldehyde
photochemically.
Marine algae were also shown to produce
photosensitizers (
12).
These facts suggest that photolysis
of DMS is also affected by
biological activities, although we are not
able to estimate the
effects
yet.
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ACKNOWLEDGMENTS |
We thank Terumi Tanimoto in Chugoku National Industrial Research
Institute for his help in collecting marine samples. We also thank
Misaki Ohta at Towa Kagaku Co., Ltd., for her help in determining the
16S ribosomal DNA sequences.
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FOOTNOTES |
*
Corresponding author. Mailing address: Chugoku National
Industrial Research Institute, 2-2-2 Hirosuehiro, Kure, Hiroshima 737-0197, Japan. Phone: 81-823-72-1936 or 81-823-72-1934. Fax: 81-823-73-3284. E-mail: fuse{at}cniri.go.jp.
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REFERENCES |
| 1.
|
Albert, D. B., and C. S. Martens.
1997.
Determination of low-molecular-weight organic acid concentrations in seawater and pore-water samples via HPLC.
Mar. Chem.
56:27-37.
|
| 2.
|
Brimblecombe, P., and D. Shooter.
1986.
Photo-oxidation of dimethylsulphide in aqueous solution.
Mar. Chem.
19:343-353[CrossRef].
|
| 3.
|
Brugger, A.,
D. Slezak,
I. Obernosterer, and G. J. Herndl.
1998.
Photolysis of dimethylsulfide in the northern Adriatic Sea: dependence on substrate concentration, irradiance and DOC concentration.
Mar. Chem.
59:321-331[CrossRef].
|
| 4.
|
Charlson, R. J.,
J. E. Lovelock,
M. O. Andreae, and S. G. Warren.
1987.
Ocean phytoplankton, atmospheric sulphur, cloud albedo and climate.
Nature
326:655-661[CrossRef].
|
| 5.
|
Cho, K. S.,
M. Hirai, and M. Shoda.
1991.
Degradation characteristics of hydrogen sulfide, methanethiol, dimethyl sulfide and dimethyl disulfide by Thiobacillus thioparus DW44 isolated from peat biofilter.
J. Ferment. Bioeng.
71:384-389[CrossRef].
|
| 6.
|
Dacey, J. H. W.,
S. G. Wakeham, and B. L. Howes.
1984.
Henry's law constants for dimethylsulfide in freshwater and seawater.
Geophys. Res. Lett.
11:202-208.
|
| 7.
|
De Bont, J. A. M.,
J. P. van Dijken, and W. Harder.
1981.
Dimethyl sulphoxide and dimethyl sulphide as a carbon, sulphur and energy source for growth of Hyphomicrobium S.
J. Gen. Microbiol.
127:315-323.
|
| 8.
|
De Marco, P.,
P. Moradas-Ferreira,
T. P. Higgins,
I. McDonald,
E. M. Kenna, and J. C. Murrell.
1999.
Molecular analysis of a novel methanesulfonic acid monooxygenase from the methylotroph Methylosulfonomonas methylovora.
J. Bacteriol.
181:2244-2251[Abstract/Free Full Text].
|
| 9.
|
De Zwart, J. M. M.,
P. N. Nelisse, and J. G. Kuenen.
1996.
Isolation and characterization of Methylophaga sulfidovorans sp. nov.: an obligately methylotrophic, aerobic, dimethylsulfide oxidizing bacterium from a microbial mat.
FEMS Microbiol. Lett.
20:261-270.
|
| 10.
| Eichhorn, E., J. R. van der Ploeg, and T. Leisinger. 1999. Characterization of a two-component
alkanesulfonate monooxygenase from Escherichia coli.
274:26639-26646.
|
| 11.
|
Fuse, H.,
M. Ohta,
O. Takimura,
K. Murakami,
H. Inoue,
Y. Yamaoka,
J. M. Oclarit, and T. Omori.
1998.
Oxidation of trichloroethylene and dimethyl sulfide by a marine Methylomicrobium strain containing soluble methane monooxygenase.
Biosci. Biotechnol. Biochem.
62:1925-1931[CrossRef][Medline].
|
| 12.
|
Fuse, H.,
O. Takimura,
K. Kamimura,
K. Murakami,
Y. Yamaoka, and Y. Murooka.
1995.
Transformation of dimethyl sulfide and related compounds by cultures and cell extracts of marine phytoplankton.
Biosci. Biotechnol. Biochem.
59:1773-1775.
|
| 13.
|
Gonzalez, J. M.,
F. Mayer,
M. A. Moran,
R. E. Hodson, and W. B. Whitman.
1997.
Microbulbifer hydrolyticus gen. nov., sp. nov., and Marinobacterium georgiense gen. nov., sp. nov., two marine bacteria from a lignin-rich pulp mill waste enrichment community.
Int. J. Syst. Bacteriol.
47:369-376[Abstract/Free Full Text].
|
| 14.
|
Gonzalez, J. M.,
R. P. Kiene, and M. A. Moran.
1999.
Transformation of sulfur compounds by an abundant lineage of marine bacteria in the -subclass of the class Proteobacteria.
Appl. Environ. Microbiol.
65:3810-3819[Abstract/Free Full Text].
|
| 15.
|
Hanlon, S. P.,
R. A. Holt,
G. R. Moore, and A. G. McEwan.
1994.
Isolation and characterization of a strain of Rhodobacter sulfidophilus: a bacterium which grows autotrophically with dimethylsulphide as electron donor.
Microbiology
140:1953-1958.
|
| 16.
|
Hatakeyama, S.,
M. Okuda, and H. Akimoto.
1982.
Formation of sulfur dioxide and methanesulfonic acid in the photooxidation of dimethyl sulfide in the air.
Geophys. Res. Lett.
9:583-586.
|
| 17.
|
Horinouchi, M.,
K. Kasuga,
H. Nojiri,
H. Yamane, and T. Omori.
1997.
Cloning and characterization of genes encoding an enzyme which oxidizes dimethyl sulfide in Acinetobacter sp. Strain 20B.
FEMS Microbiol. Lett.
155:99-105[CrossRef][Medline].
|
| 18.
|
Juliette, L. Y.,
M. R. Hyman, and D. J. Arp.
1993.
Inhibition of ammonia oxidation in Nitrosomonas europaea by sulfur compounds: thioethers are oxidized to sulfoxides by ammonia monooxygenase.
Appl. Environ. Microbiol.
59:3718-3727[Abstract/Free Full Text].
|
| 19.
|
Kanagawa, T., and D. P. Kelly.
1986.
Breakdown of dimethyl sulphide by mixed cultures and by Thiobacillus thioparus.
FEMS Microbiol. Lett.
34:13-19[CrossRef].
|
| 20.
|
Keller, M. D.,
W. K. Bellows, and R. R. L. Guillard.
1989.
Dimethyl sulfide production in marine phytoplankton, p. 167-182.
In
E. S. Saltzman, and W. J. Cooper (ed.), Biogenic sulfur in the environment. American Chemical Society, Washington, D.C.
|
| 21.
|
Kieber, D. J.,
J. Jiao,
R. P. Kiene, and T. S. Bates.
1996.
Impact of dimethylsulfide photochemistry on methyl sulfur cycling in equatorial Pacific Ocean.
J. Geophys. Res.
101:3715-3722[CrossRef].
|
| 22.
|
Kieber, R. J., and K. Mopper.
1990.
Determination of picomolar concentration of carbonyl compounds in natural water, including seawater, by liquid chromatography.
Environ. Sci. Technol.
24:1477-1481[CrossRef].
|
| 23.
|
Kiene, R. P., and T. S. Bates.
1990.
Biological removal of dimethyl sulfide from sea water.
Nature
345:702-705[CrossRef].
|
| 24.
|
Kiene, R. P.,
L. J. Linn,
J. González,
M. A. Moran, and J. A. Bruton.
1999.
Dimethylsulfoniopropionate and methanethiol are important precursors of methionine and protein-sulfur in marine bacterioplankton.
Appl. Environ. Microbiol.
65:4549-4558[Abstract/Free Full Text].
|
| 25.
|
Montoya, G.,
C. Svensson,
H. Savage,
J. D. Schwenn, and I. Sinning.
1998.
Crystallization and preliminary X-ray diffraction studies of phospho-adenylylsulfate (PAPS) reductase from E. coli.
Acta Crystallogr. D.
54:281-283[CrossRef][Medline].
|
| 26.
|
Omori, T.,
Y. Saiki,
K. Kasuga, and T. Kodama.
1995.
Desulfurization of alkyl and aromatic sulfides and sulfonates by dibenzothiophene-desulfurizing Phodococcus sp. strain SY1.
Biosci. Biotechnol. Biochem.
59:1195-1198.
|
| 27.
|
Oppenheimer, C. H., and C. E. ZoBell.
1952.
The growth and viability of sixty-three species of marine bacteria as influenced by hydrostatic pressure.
J. Mar. Res.
11:10-18.
|
| 28.
|
Schwartz, S. E.
1988.
Are global cloud albedo and climate controlled by marine phytoplankton?
Nature
336:441-445[CrossRef].
|
| 29.
|
Smith, N. A., and D. P. Kelly.
1988.
Isolation and physiological characterization of autotrophic sulphur bacteria oxidizing dimethyl disulphide as sole source of energy.
J. Gen. Microbiol.
134:1407-1417.
|
| 30.
|
Suylen, G. M. H., and J. G. Kuenen.
1986.
Chemostat enrichment and isolation of Hyphomicrobium EG.
Antonie Leeuwenhoek
52:281-293.
|
| 31.
|
Visscher, P. T., and H. van Gemerden.
1991.
Photo-autotrophic growth of Thiocapsa roseopersicina on dimethyl sulfide.
FEMS Microbiol. Lett.
81:247-250[CrossRef].
|
| 32.
|
Visscher, P. T., and B. F. Taylor.
1993.
A new mechanism for the aerobic catabolism of dimethyl sulfide.
Appl. Environ. Microbiol.
59:3784-3789[Abstract/Free Full Text].
|
| 33.
|
Visscher, P. T., and B. F. Taylor.
1993.
Aerobic and anaerobic degradation of a range of alkyl sulfides by a denitrifying marine bacterium.
Appl. Environ. Microbiol.
59:4083-4089[Abstract/Free Full Text].
|
| 34.
|
Weiner, J. H.,
D. P. MacIsaac,
R. E. Bishop, and P. T. Bilous.
1988.
Purification and properties of Escherichia coli dimethyl sulfoxide reductase, an iron-sulfur molybdoenzyme with broad substrate specificity.
J. Bacteriol.
170:1505-1510[Abstract/Free Full Text].
|
| 35.
|
Yin, F.,
D. Grosjean,
R. C. Flagan, and J. H. Seinfeld.
1990.
Photooxidation of dimethyl sulfide and dimethyl disulfide. II. Mechanism evaluation.
J. Atmos. Chem.
11:309-364[CrossRef].
|
| 36.
|
Zeyer, J.,
P. Eicher,
S. G. Wakeham, and R. P. Schwarzenbach.
1987.
Oxidation of dimethyl sulfide to dimethyl sulfoxide by phototrophic purple bacteria.
Appl. Environ. Microbiol.
53:2026-2032[Abstract/Free Full Text].
|
| 37.
|
Zhang, L.,
I. Kuniyoshi,
M. Hirai, and M. Shoda.
1991.
Oxidation of dimethyl sulfide by Pseudomonas acidovorans DMR-11 isolated from peat biofilter.
Biotechnol. Lett.
13:223-228.
|
| 38.
|
Zhang, L.,
M. Hirai, and M. Shoda.
1991.
Removal characteristics of dimethyl sulfide, methanethiol and hydrogen sulfide by Hyphomicrobium sp. I55 isolated from peat biofilter.
J. Ferment. Bioeng.
72:392-396.
|
Applied and Environmental Microbiology, December 2000, p. 5527-5532, Vol. 66, No. 12
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Copyright © 2000, American Society for Microbiology. All rights reserved.
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