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Appl Environ Microbiol, April 1998, p. 1576-1579, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Anaerobic Degradation of Hexadecan-2-one by a
Microbial Enrichment Culture under Sulfate-Reducing
Conditions
Agnes
Hirschler,1,*
Jean-Francois
Rontani,2
Danielle
Raphel,2
Robert
Matheron,1 and
Jean-Claude
Bertrand2
Laboratoire de Microbiologie, Service 452,
Faculté des Sciences et Techniques de
Saint-Jérôme, 13397 Marseille Cedex
20,1 and
Laboratoire
d'Océanographie et de Biogéochimie UMR 6535, Centre
d'Océanologie de Marseille OSU, Faculté des Sciences
de Luminy, 13288 Marseille,2 France
Received 14 July 1997/Accepted 22 January 1998
 |
ABSTRACT |
A microbial enrichment culture from marine sediment was able to
grow on hexadecan-2-one as the sole source of carbon and energy under
sulfate-reducing conditions. Oxidation of the ketone involved carboxylation reactions and was coupled to sulfide production. This
enrichment culture also grew on 6,10,14-trimethylpentadecan-2-one.
| |
TEXT |
Degradation of hydrocarbons under
anoxic conditions has been demonstrated in microbial communities and
pure cultures of sulfate-reducing (1, 2, 7, 15, 18, 26, 31),
denitrifying (5, 17, 27), and iron(III)-reducing bacteria
(19). Most of these studies involved aromatic hydrocarbons.
Indeed, there are only a few reports on the anaerobic degradation of
aliphatic hydrocarbons, particularly alkanes (1, 5, 20, 31).
Due to the lack of unsaturations in alkanes, their degradation is
difficult in the absence of molecular oxygen.
While alkanes cannot be directly photooxidized by solar light, there
are several compounds in seawater (10) or petroleum (29) which are capable of introducing oxygen atoms into
these hydrocarbon molecules by photochemical mechanisms. The
photosensitized oxidation of n-alkanes generates mainly
ketones and secondary alcohols with the same number of carbon atoms as
the alkanes (11, 14), while oxidation of branched alkanes
leads mainly to the formation of tertiary alcohols and ketones
(30).
When photooxidation products are not fully degraded under aerobic
conditions, they will reach anoxic environments where they might be
degraded. Further, in the literature, there are relatively few reports
dealing with degradation under anaerobic conditions of ketones and
secondary alcohols resulting from hydrocarbon photooxidation. Acetone
degradation has been shown to occur in an enrichment culture under
methanogenic conditions (23) and also in pure cultures using
nitrate (3, 24) or sulfate (25) as the electron
acceptor. Higher ketones, such as butanone, pentan-2-one, hexan-2-one,
and hexan-3-one, were degraded anaerobically by pure cultures of
denitrifying bacteria (24). A pure culture of
sulfate-reducing bacteria was also able to degrade butanone
(13). A recent study showed the degradation of a branched
isoprenoid ketone, 6,10,14-trimethylpentadecan-2-one, by a denitrifying
marine bacterium (28). However, no evidence for microbial
degradation of ketones higher than butanone or secondary alcohols under
sulfate-reducing conditions has been presented. Owing to the abundance
of sulfate in anoxic marine environments, experiments were conducted
under sulfate-reducing conditions with hexadecan-2-one,
nonadecan-10-one, and two pristane photo-oxidation products
(6,10,14-trimethylpentadecan-2-one and
2,6,10,14-tetramethylpentadecan-2-ol).
Two of the compounds tested, hexadecan-2-one and
6,10,14-trimethylpentadecan-2-one, were degraded under sulfate-reducing
conditions in microcosms and enrichment cultures.
Chemicals.
Hexadecan-2-one and nonadecan-10-one were purchased
from Aldrich (Milwaukee, Wis.). 6,10,14-Trimethylpentadecan-2-one
was produced by oxidation of phytol (Riedel de Haën) with
KMnO4 in acetone (6).
2,6,10,14-Tetramethylpentadecan-2-ol was obtained by condensation of
6,10,14-trimethylpentadecan-2-one and methylmagnesium iodide
in anhydrous diethyl ether. Reduction of
6,10,14-trimethylpentadecan-2-one with LiAlH4 in diethyl
ether gave the corresponding secondary alcohol. The synthesis of
2-(1-hydroxyethyl)pentadecanoic acid required three steps: (i)
condensation of ethyl acetoacetate with 1-bromotridecane in the
presence of potassium tert-butoxide (12), (ii)
reduction of the resulting 
-ketoester with NaBH4
(which did not reduce the carboxyl group of esters) for 15 min in
methanol, and (iii) alkaline hydrolysis of the produced
-hydroxyester (5% KOH in 50% CH3OH, reflux for 1 h).
Sediment incubation protocol.
Microcosms were prepared in
60-ml serum bottles from a volume of sediment and a volume of
sterilized site water enriched as described below. Coastal sediment
showing little oil contamination (0.06 kg/kg of sediment) was collected
at Fos Bay (France). Site water was enriched with 3.7 mM ammonium
chloride, 2 mM sodium thiosulfate, 14 mM morpholinepropanesulfonic acid
(pH 7.2), and 1 mg of resazurin per liter. The fresh sediment and water
were homogenized under an N2-H2-CO2
(85:10:5) atmosphere in an anaerobic chamber (La Calhène). A
mixture of four compounds, hexadecan-2-one, nonadecan-10-one,
6,10,14-trimethylpentadecan-2-one, and
2,6,10,14-tetramethylpentadecan-2-ol, had previously been added
to the empty bottles; the compounds were dissolved in dichloromethane
and distributed so as to provide a final concentration of 200 mg/liter
in each microcosm. Dichloromethane was then allowed to evaporate from
the serum bottles. Killed controls were prepared with autoclaved
sediments. The bottles were sealed with Teflon-coated butyl rubber
stoppers (Serflam, Marseille, France) and incubated at 30°C in the
dark.
Subculture procedure.
A homogeneous fraction (3 ml) of the
microcosm was transferred to serum bottles containing the substrate
hexadecan-2-one at a final concentration of 200 mg/liter in artificial
seawater medium (27 ml) with 0.4 mM Na2S · 9H2O. Hexadecan-2-one was added as described above and
autoclaved under an N2-CO2 (90:10) atmosphere. The seawater medium contained (per liter, unless otherwise noted) 23.5 g of NaCl, 10.6 g of MgCl2 · 6H2O, 3.9 g of Na2SO4,
0.19 g of NaHCO3, 0.66 g of KCl, 0.1 g of
KBr, 0.024 g of H3BO3, 0.040 g of
SrCl2 · 6H2O, 0.2 g of
NH4Cl, 50 mg of yeast extract, 1 mg of resazurin, and
3 g of morpholinepropanesulfonic acid (pH 7.2). After autoclaving,
10 ml of CaCl2 · 2H2O (140 g/liter),
1 ml of trace element solution SL12 (21), 1 ml of
selenite-tungstate solution (33), 1 ml of 7-vitamin solution
(22), and 4 ml of phosphate buffer (4.4 g of
NaH2PO4 · 2H2O and 25.8 g of Na2HPO4 · 12H2O per
liter) were added from sterile stock solutions. Prior to
inoculation, the medium was incubated overnight in an
anaerobic chamber. The bottles were incubated at 30°C in the dark on
a reciprocal shaker (100 strokes/min).
Analytical procedures.
Sulfide was quantified with
Cu2+ ions yielding CuS (8).
For analyses of the ketones and alcohol, the sediment and the aqueous
phase from the microcosm were separated by decantation. The wet
sediment was extracted under sonication with isopropanol-hexane (4:1,
vol/vol) (9), and the aqueous phase was extracted three times with chloroform. For analysis of the substrate of the subculture without sediment, the entire contents of the vial were extracted with
chloroform. The combined hexane and chloroform extracts were dried with
anhydrous Na2SO4, filtered, and concentrated by
means of rotary evaporation to give extract E1 (containing residual substrates and eventual nonacidic metabolites). The aqueous phase was
then acidified with hydrochloric acid (pH 1) and extracted three times
with chloroform to give extract E2 (containing acidic compounds). After
evaporation of the solvents, extracts E1 and E2 were taken up
separately in 250 µl of a mixture of pyridine and
N,O-bistrimethylsilyl-trifluoroacetamide
(3:1, vol/vol) and allowed to silylate at 50°C for 1 h.
Following evaporation to dryness under nitrogen, the residues were
dissolved in hexane and analyzed by gas chromatography-mass
spectrometry (GC/MS). Compounds were identified (by comparison of their
retention times and mass spectra with those of standards) and
quantified (after calibration with external standards) by GC/MS. The
GC/MS analyses were carried out with an HP 5890 series II plus gas
chromatograph connected to an HP 5972 mass spectrometer. The operating
conditions included a capillary measuring 15 m by 0.25 mm (inside
diameter), a column coated with BPX 35 (SGE), an oven temperature
programmed to go from 60 to 150°C at 30°C/min and then from 150 to
320°C at 3°C/min, a carrier gas (He) pressure of 0.48 bar, an
injector temperature of 320°C, an electron energy of 70 eV, and a
source temperature of 170°C.
The microbial biomass was estimated indirectly by quantifying the
cellular protein (4) assuming a 50% protein content.
Depletion experiments in microcosms.
In microcosms fed with
the mixture of four compounds, hexadecan-2-one was the first compound
to be depleted. Within 2 months, more than 80% was depleted while only
15% of the 6,10,14-trimethylpentadecan-2-one was depleted. After 10 more months, 42% of the 6,10,14-trimethylpentadecan-2-one was depleted
with respect to a control incubated just as long. Neither
nonadecan-10-one nor 2,6,10,14-tetramethylpentadecan-2-ol was depleted
after 1 year of incubation. In the controls, none of the compounds
disappeared within 6 months; however, after 1 year of incubation, only
80% of the four substrates remained. Successive subcultures confirmed
those depletions. We conclude that of the compounds studied,
hexadecan-2-one is the most rapidly depleted.
Evidence that hexadecan-2-one metabolism is linked to sulfate
reduction.
Successive transfers to defined medium with
hexadecan-2-one were made to establish enrichment cultures
without sediment. To link sulfate reduction and
hexadecan-2-one oxidation, transfers were made in complete
artificial seawater medium and in sulfate-depleted artificial seawater
medium. After 1 month of incubation, 80% of the hexadecan-2-one
was degraded in the complete artificial seawater medium while only 10%
was degraded in the sulfate-depleted artificial seawater medium.
Transfer of the enrichment culture to medium with hexadecan-2-one
and/or yeast extract showed that yeast extract alone in artificial
seawater medium did not support growth, while growth was the same on
hexadecan-2-one with or without yeast extract. On the other hand, the
weak growth in subcultures without sulfate was detectable only in the
presence of both hexadecan-2-one and yeast extract. In conclusion, the
experiments show that efficient anaerobic degradation of
hexadecan-2-one requires sulfate and the activity of sulfate-reducing
bacteria. Nevertheless, some degradation occurred in the absence of
sulfate, and hence, other microorganisms are probably involved.
The community growing in yeast extract-free artificial seawater medium
with hexadecan-2-one as the sole source of carbon was
composed of
curved rods and of some oval and rod-shaped cells
(Fig.
1). The culture grown on hexadecan-2-one
as the sole source
of carbon and energy reached its optimum growth
within about 20
days with an optical density at 450 nm of about 0.20. The curve
is not shown, as the culture often formed aggregates or
biofilm
on a glass surface. Nevertheless, increased optical density was
linked to an increase in microbial biomass, as about 16 mg (dry
weight)
per liter was produced in a month of growth while 0.6
mM
hexadecan-2-one was consumed. At optimum growth, 60% of the
hexadecan-2-one was degraded. Although sulfate was not limiting,
about
35% of the hexadecan-2-one remained at the end of the experiment
(Fig.
2A). The degradation of
hexadecan-2-one was linked to sulfide
production (Fig.
2A and B).
In sterile controls, no depletion
occurred and no sulfide was produced.
Furthermore, in inoculated
controls without substrate, the sulfide
produced never exceeded
0.2 mM. At the end of the experiment,
5.42 mM sulfide was produced
during the degradation of 0.53 mM
hexadecan-2-one. The stoichiometry
of complete
hexadecan-2-one oxidation through sulfate reduction
is as
follows:
CH
3(CH
2)
13COCH
3 + 11.75 SO
42
11.75 S
2 + 16 CO
2 + 16 H
2O. Therefore, the amount of sulfide
produced
will allow the oxidation of 0.46 mM hexadecan-2-one. The
remaining
0.07 mM will be converted to cell carbon, and possibly to
energy,
by the nonsulfidogenic part of the community. Since it was
consistent
with the cell mass produced (0.04 mM hexadecanone turned
into
cell carbon), we concluded that oxidation was complete.
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FIG. 1.
Phase-contrast photomicrograph of an enrichment culture
grown on hexadecan-2-one and sulfate. Bar, 10 µm.
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FIG. 2.
Concentrations of hexadecan-2-one (A) and sulfide (B) in
an enrichment culture growing in the presence of hexadecan-2-one. Open
bars correspond to sterile controls. The data are mean values from
three cultures. At each time, the entire contents of three vials were
analyzed.
|
|
After growth of the enrichment culture on hexadecan-2-one, we detected
2-(1-hydroxyethyl)pentadecanoic acid in extract E2
(Fig.
3A). This metabolite, which was
undetectable in controls
with a sterilized inoculum, was identified
unambiguously by comparison
of its retention time and mass spectrum
(Fig.
3B) with those of
a synthesized standard. The presence of this
compound clearly
shows that the metabolism of hexadecan-2-one by the
enrichment
culture involves carboxylation reactions. Such a mechanism
has
already been described for the anaerobic oxidation of acetone
(
3,
23,
24). Partial reduction of the carbonyl group to
the
corresponding secondary alcohol was also observed. The involvement
of
this "blind-alley" pathway probably results from nonspecific
enzyme
activity not related to hexadecan-2-one degradation (
16).
On
the basis of these results, we propose the pathway described
in the
Fig.
4 for the metabolism of
hexadecan-2-one by the enrichment
culture. The detection of a
relatively strong proportion of pentadecanoic
acid in extract E2 (Fig.
3A) is in good agreement with such a
pathway. If carboxylation of
ketones is easier than that of the
corresponding alcohols, due to
activation of the carbon
relative
to the carbonyl group by
keto-enol tautomerism, the possibility
of hexadecan-2-ol carboxylation
cannot be completely excluded
(
3,
32).
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FIG. 3.
(A) Total ion chromatogram of (silylated) extract E2
obtained after the growth of an enrichment culture on hexadecan-2-one.
(B) Electron impact mass spectrum of metabolite 8, identified as
(silylated) 2-(1-hydroxyethyl)pentadecanoic acid.
|
|
Although the carboxylation takes place on the secondary carbon in the
position relative to the carbonyl group of hexadecan-2-one,
the
enrichment culture was unable to degrade nonadecan-10-one.
An attempt
at hexadecan-3-one biodegradation confirmed that only
ketones with the
keto group at position 2 could be degraded by
the enrichment culture.
These results confirm once more the importance of the community of
sulfate-reducing bacteria in anoxic environments and, in
particular, in
the degradation of hydrocarbons. Furthermore, it
is the first
demonstration of the degradation of a typical hydrocarbon
photooxidation product under sulfate-reducing conditions.
| |
ACKNOWLEDGMENTS |
We are indebted to F. Widdel for helpful discussions and to
R. Guyoneaud for microphotograph preparation.
This work was supported by a grant from the Centre National de la
Recherche Scientifique and Elf Aquitaine through Research Groupe HYCAR
1123.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Laboratoire de
Microbiologie, Service 452, Faculté des Sciences et Techniques de
Saint-Jérôme, 13397 Marseille Cedex 20, France. Phone: 04 91 28 81 91. Fax: 04 91 28 80 30. E-mail:
agnes.hirschler{at}microbio.u-3mrs.fr.
| |
REFERENCES |
| 1.
|
Aeckersberg, F.,
F. Bak, and F. Widdel.
1991.
Anaerobic oxidation of saturated hydrocarbons to CO2 by a new type of sulfate-reducing bacterium.
Arch. Microbiol.
156:5-14.
|
| 2.
|
Beller, H. R.,
A. M. Spormann,
P. K. Sharma,
J. R. Cole, and M. Reinhard.
1996.
Isolation and characterization of a novel toluene-degrading, sulfate-reducing bacterium.
Appl. Environ. Microbiol.
62:1188-1196[Abstract].
|
| 3.
|
Bonnet-Smits, E. M.,
L. A. Robertson,
J. P. Van Dijken,
E. Senior, and J. G. Kuenen.
1988.
Carbon dioxide fixation as the initial step in the metabolism of acetone by Thiosphaera pantotropha.
J. Gen. Microbiol.
134:2281-2289.
|
| 4.
|
Bradford, M. M.
1976.
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem.
72:248-254[Medline].
|
| 5.
|
Bregnard, T. P. A.,
P. Höhener,
A. Häner, and J. Zeyer.
1996.
Degradation of weathered diesel fuel by microorganisms from a contaminated aquifer in aerobic and anaerobic microcosms.
Environ. Toxicol. Chem.
15:299-307.
|
| 6.
|
Cason, J., and D. W. Graham.
1965.
Isolation of isoprenoid acids from a California petroleum.
Tetrahedron
21:471-483.
|
| 7.
|
Coates, J. D.,
R. T. Anderson, and D. R. Lovley.
1996.
Oxidation of polycyclic aromatic hydrocarbons under sulfate-reducing conditions.
Appl. Environ. Microbiol.
62:1099-1101[Abstract].
|
| 8.
|
Cord-Ruwisch, R.
1985.
A quick method for the determination of dissolved and precipitated sulfides in cultures of sulfate-reducing bacteria.
J. Microbiol. Methods
4:33-36.
|
| 9.
|
de Leeuw, J. W.,
B. R. T. Simoneit,
J. J. Boon,
W. I. C. Rijpstra,
F. de Lange,
J. C. W. van der Linden,
V. A. Correia,
A. L. Burlingame, and P. A. Schenck.
1977.
Phytol derived compounds in the geosphere, p. 61-79. In
R. Campos, and J. Goni (ed.), Advances in organic geochemistry 1975.
Enadimsa, Madrid, Spain.
|
| 10.
|
Ehrhardt, M.,
F. Bouchertall, and P. Hopf.
1982.
Aromatic ketones concentrated from Baltic Sea water.
Mar. Chem.
11:449-461.
|
| 11.
|
Ehrhardt, M., and G. Petrick.
1985.
The sensitized photo-oxidation of n-pentadecane as a model for abiotic decomposition of aliphatic hydrocarbons in seawater.
Mar. Chem.
16:227-238.
|
| 12.
|
Etheredge, S. J.
1966.
Bicyclic ketones by intramolecular alkylation: a reinvestigation.
J. Am. Chem. Soc.
12:1990-1994.
|
| 13.
|
Galushko, A. S., and E. P. Rozanova.
1992.
Desulfobacterium cetonicum sp. nov.: a sulfate-reducing bacterium which oxidizes fatty acids and ketones.
Microbiology
60:742-746. (English translation from Mikrobiologiya 60:102-107, 1991.)
|
| 14.
|
Guiliano, M.,
F. El Anba-Lurot,
P. Doumenq,
G. Mille, and J. F. Rontani.
1997.
Photo-oxidation of n-alkanes in simulated marine environmental conditions.
J. Photochem. Photobiol. Ser. A
102:127-132.
|
| 15.
|
Haag, F.,
M. Reinhard, and P. L. McCarty.
1991.
Degradation of toluene and p-xylene in anaerobic microcosms: evidence for sulfate as a terminal electron acceptor.
Environ. Toxicol. Chem.
10:1379-1389.
|
| 16.
|
Klug, M. J., and A. J. Markovetz.
1971.
Utilization of aliphatic hydrocarbons by microorganisms.
Adv. Microb. Physiol.
5:1-43[Medline].
|
| 17.
|
Kuhn, E. P.,
J. Zeyer,
P. Eicher, and R. P. Schwarzenbach.
1988.
Anaerobic degradation of alkylated benzenes in denitrifying laboratory aquifer columns.
Appl. Environ. Microbiol.
54:490-496[Abstract/Free Full Text].
|
| 18.
|
Lovley, D. R.,
J. D. Coates,
J. C. Woodward, and E. J. P. Phillips.
1995.
Benzene oxidation coupled to sulfate reduction.
Appl. Environ. Microbiol.
61:953-958[Abstract].
|
| 19.
|
Lovley, D. R., and D. J. Lonergan.
1990.
Anaerobic oxidation of toluene, phenol, and p-cresol by the dissimilatory iron-reducing organism, GS-15.
Appl. Environ. Microbiol.
56:1858-1864[Abstract/Free Full Text].
|
| 20.
|
Morikawa, M.,
M. Kanemoto, and T. Imanaka.
1996.
Biological oxidation of alkane to alkene under anaerobic conditions.
J. Ferment. Bioeng.
82:309-311.
|
| 21.
|
Pfennig, N., and H. G. Trüper.
1992.
The family Chromatiaceae, p. 3200-3221. In
A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K. H. Schleifer (ed.), The prokaryotes, 2nd ed.
Springer-Verlag, New York, N.Y.
|
| 22.
|
Pfennig, N.,
F. Widdel, and H. G. Trüper.
1981.
The dissimilatory sulfate-reducing bacteria, p. 926-940. In
M. P. Starr, H. Stolp, H. G. Trüper, A. Balows, and H. G. Schlegel (ed.), The prokaryotes, 1st ed.
Springer-Verlag, New York, N.Y.
|
| 23.
|
Platen, H., and B. Schink.
1987.
Methanogenic degradation of acetone by an enrichment culture.
Arch. Microbiol.
149:136-141[Medline].
|
| 24.
|
Platen, H., and B. Schink.
1989.
Anaerobic degradation of acetone and higher ketones via carboxylation by newly isolated denitrifying bacteria.
J. Gen. Microbiol.
135:883-891[Abstract/Free Full Text].
|
| 25.
|
Platen, H.,
A. Temmes, and B. Schink.
1990.
Anaerobic degradation of acetone by Desulfococcus biacutus spec. nov.
Arch. Microbiol.
154:355-361[Medline].
|
| 26.
|
Rabus, R.,
M. Fukui,
H. Wilkes, and F. Widdel.
1996.
Degradative capacities and 16S rRNA-targeted whole-cell hybridization of sulfate-reducing bacteria in an anaerobic enrichment culture utilizing alkylbenzenes from crude oil.
Appl. Environ. Microbiol.
62:3605-3613[Abstract].
|
| 27.
|
Rabus, R., and F. Widdel.
1996.
Utilization of alkylbenzenes during anaerobic growth of pure cultures of denitrifying bacteria on crude oil.
Appl. Environ. Microbiol.
62:1238-1241[Abstract].
|
| 28.
|
Rontani, J.-F.,
M. J. Gilewicz,
V. D. Michotey,
T. L. Zheng,
P. C. Bonin, and J.-C. Bertrand.
1997.
Aerobic and anaerobic metabolism of 6,10,14-trimethylpentadecan-2-one by a denitrifying bacterium isolated from marine sediments.
Appl. Environ. Microbiol.
63:636-643[Abstract].
|
| 29.
|
Rontani, J.-F., and P. J.-P. Giral.
1990.
Significance of photochemical degradation of petroleum hydrocarbon fractions in seawater.
Int. J. Environ. Anal. Chem.
42:61-68.
|
| 30.
|
Rontani, J.-F., and G. Giusti.
1987.
Photosensitized oxidation of pristane in sea water: effect of photochemical reactions on tertiary carbons.
J. Photochem. Photobiol. Ser. A
40:107-120.
|
| 31.
|
Rueter, P.,
R. Rabus,
H. Wilkes,
F. Aeckersberg,
F. A. Rainey,
H. W. Jannasch, and F. Widdel.
1994.
Anaerobic oxidation of hydrocarbons in crude oil by new types of sulphate-reducing bacteria.
Nature (London)
372:455-458[Medline].
|
| 32.
|
Schink, B.
1988.
Principles and limits of anaerobic degradation: environmental and technological aspects, p. 771-846. In
A. J. B. Zehnder (ed.), Biology of anaerobic microorganisms.
Wiley, New York, N.Y.
|
| 33.
|
Widdel, F., and F. Bak.
1992.
Gram-negative mesophilic sulfate-reducing bacteria, p. 3353-3378. In
A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K. H. Schleifer (ed.), The prokaryotes, 2nd ed.
Springer-Verlag, New York, N.Y.
|
Appl Environ Microbiol, April 1998, p. 1576-1579, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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