APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Sept. 1998, p. 3507–3511
0099-2240/98/$04.0010
Copyright © 1998, American Society for Microbiology. All Rights Reserved.
Vol. 64, No. 9
Pseudomonas sp. Strain 273, an Aerobic a,v-DichloroalkaneDegrading Bacterium
CATRIN WISCHNAK,1 FRANK E. LÖFFLER,1,2 JIERAN LI,2 JOHN W. URBANCE,2
1
AND RUDOLF MÜLLER *
Arbeitsbereich Biotechnologie II, Technische Universität Hamburg-Harburg, D-21071 Hamburg,
Germany,1 and Center for Microbial Ecology, Michigan State University,
East Lansing, Michigan 48824-13252
Received 16 March 1998/Accepted 1 July 1998
A gram-negative, aerobic bacterium was isolated from soil; this bacterium grew in 50% (vol/vol) suspensions
of 1,10-dichlorodecane (1,10-DCD) as the sole source of carbon and energy. Phenotypic and small-subunit ribosomal RNA characterizations identified the organism, designated strain 273, as a member of the genus Pseudomonas. After induction with 1,10-DCD, Pseudomonas sp. strain 273 released stoichiometric amounts of chloride
from C5 to C12 a,v-dichloroalkanes in the presence of oxygen. No dehalogenation occurred under anaerobic
conditions. The best substrates for dehalogenation and growth were C9 to C12 chloroalkanes. The isolate also
grew with nonhalogenated aliphatic compounds, and decane-grown cells dechlorinated 1,10-DCD without a lag
phase. In addition, cells grown on decane dechlorinated 1,10-DCD in the presence of chloramphenicol, indicating that the 1,10-DCD-dechlorinating enzyme system was also induced by decane. Other known alkane-degrading Pseudomonas species did not grow with 1,10-DCD as a carbon source. Dechlorination of 1,10-DCD was
demonstrated in cell extracts of Pseudomonas sp. strain 273. Cell-free activity was strictly oxygen dependent,
and NADH stimulated dechlorination, whereas EDTA had an inhibitory effect.
that grew aerobically on C7 and C9 chloro-, bromo-, and iodoalkanes. Another Pseudomonas species with an n-alkane-inducible enzyme system dehalogenated these compounds cometabolically when grown on n-alkanes (19). The chlorinated
analogues did not induce the oxygenolytic enzyme system,
which might explain why it is difficult to enrich for chloroalkane degraders.
We monitored various soil samples from pristine and contaminated sites for their potential to release chloride ions from
1,10-DCD over 5 years. This article described the isolation and
characterization of a novel Pseudomonas species that was able
to grow in suspensions of 1,10-DCD and other a,v-dichloroalkanes.
Garden soil samples (5 g [dry weight]) from five different locations in the area of Stuttgart, Germany, were suspended in
50-ml portions of mineral salts medium. Mineral salts medium
contained the following (per liter): 5.37 g of Na2HPO4 z 12H2O,
1.36 g of KH2PO4, 0.5 g of (NH4)2SO4, 0.2 g of MgSO4 z 7H2O,
0.2 ml of vitamin solution, 1 ml of trace element solution,
0.05% (wt/vol) yeast extract, and 10 mmol of 1,10-DCD. The
trace element solution contained the following (per liter):
20.0 g of NaOH, 10 g of MgSO4 z 7H2O, 4 g of ZnSO4 z 7H2O,
1 g of CuSO4 z 5H2O, 3.2 g of MnSO4 z H2O, 20 g of Fe2
(SO4) z 7H2O, 100 g of Na2SO4, 1 g of NaMoO4 z 2H2O, 1 ml
of H2SO4 (concentrated), and 120 g of EDTA. The vitamin
solution contained (per liter) 100 mg of vitamin B12, 20.0 mg of
pyridoxal, 100 mg of riboflavin, and 100 mg of thiamine.
Cultures were incubated at 30°C with 10 mM 1,10-DCD as
the sole carbon and energy source and shaken at 200 rpm.
Samples (1 ml) were withdrawn weekly and analyzed for chloride release with the chlorocounter (Marius Instrumenten,
Nieuwegein, The Netherlands). Enriched cultures that had released more than 15 mM chloride were transferred (10% inoculum [vol/vol]) to fresh medium containing 10 mM 1,10DCD. Of the five different soils incubated with 1,10-DCD as
the only available carbon and energy source, chloride release
was detected from only one garden soil after 1 month. Dechlo-
Chloroparaffins are produced by chlorination of alkanes and
are classified into three groups according to their chain length:
short (C10 to C13), medium (C14 to C17), and long (C.17) chains.
They are widely used as plasticizers in polyvinylchloride, as lubricants, in paints, and in fire retardants. Their total world
production is about 300,000 metric tons per year (10). In Germany, 5,000 tons of short-chain chloroparaffins were produced
in 1994 and were mainly used in the metal industry (13). This
widespread use of chloroparaffins has raised concern and interest in the environmental fate of these water-insoluble compounds which are classified as nonbiodegradable (3).
The degradation of haloalkanes can proceed through different pathways. Haloparaffins (C12 to C18) have been reported to
be incorporated into fatty acids in bacteria, yeasts, and fungi (9,
17), resulting in their accumulation in the food chain. Another
pathway is the oxygenation at the nonhalogenated end of monohalogenated alkanes by an inherent oxygenase with a tight
substrate selectivity (5). In this case fluoroalkanes were defluorinated, but no dehalogenation was observed with chloro-,
bromo-, or iodoalkanes. Chain length was reported to have
minor effects on this oxygenation reaction. In general, aand a,v-chlorinated haloalkanes with short carbon chains
(C1 to C6) are dehalogenated hydrolytically or by a glutathione-dependent mechanism (6, 26). In contrast, a- and
a,v-haloalkanes with longer chains, e.g., 1,9-dichlorononane
and 1,10-dichlorodecane (1,10-DCD), have been proposed to
be dehalogenated by oxidative mechanisms (1, 18, 19). Studies
on the biodegradation of this class of compounds are rare,
because haloalkane-degrading microorganisms are not easily
found. Omori and Alexander (18) obtained only two bacterial
cultures from 500 enriched cultures growing on 1,9-dichlorononane. One of these organisms was a Pseudomonas species
* Corresponding author. Mailing address: Arbeitsbereich Biotechnologie II, Technische Universität Hamburg-Harburg, Denickestrasse
15, D-21071 Hamburg, Germany. Phone: 49-40-7718-3118. Fax: 49-407718-2127. E-mail: [email protected].
3507
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WISCHNAK ET AL.
rinating activity was maintained during six subsequent transfers
with 1,10-DCD as the only growth substrate. Aliquots (100 ml)
of the culture fluid were then spread onto solid mineral salts
medium with 1,10-DCD as the sole source of carbon and energy. Small, flat, opaque colonies formed within 1 week. When
streaked on HNB (5.0 g of yeast extract [Merck], 8.0 g of
nutrient broth [Difco Laboratories, Detroit, Mich.], 5.0 g of
NaCl [all per liter]) agar plates, uniform colonies were obtained. Colony material appeared homogeneous when examined by phase-contrast microscopy. BOX-PCR (targeted to
repetitive intergenic sequence elements of Streptoccus) and
ERIC-PCR (targeted to enterobacterial repetitive intergenic
consensus sequence elements) (21) performed with whole cells
grown under different conditions (HNB, tryptic soy agar, or
mineral salts medium with 1,10-DCD) gave identical banding
patterns, confirming the purity of the isolate, which was designated strain 273.
Identification of the isolate was initially performed with API
20 NE test strips (bioMérieux, Nürtingen, Germany). Further
physiological characterization of the isolate was done by the
method of Süßmuth et al. (27). The isolate was a gram-negative, motile rod, 0.8 to 1.5 mm long, with more than one polar
flagellum. Strain 273 lysed in 3% potassium hydroxide and was
aminopeptidase positive. The isolate grew aerobically and under denitrifying conditions. Nitrate was reduced to nitrite and
dinitrogen gas. No fermentative growth was observed under
anaerobic conditions. Oxidase and catalase were present. The
round sticky colonies were white to yellowish and produced a
diffusible fluorescent pigment on HNB agar plates. No polyphosphates or polyhydroxyalkanoates were formed as storage
products. Arginine dihydrolase was present. Polyamine analysis (2) revealed 9.1 mmol of spermidine, 6.8 mmol of putrescine, and 1.0 mmol of 1,3-diaminopropane per g (dry weight).
Strain 273 grew on D-glucose, gluconate, caprate, adipate, malate,
citrate, and phenylacetate. Arabinose, mannose, mannitol, Nacetylglucosamine, and maltose did not support growth. Urease, a-glucosidase, gelatin protease, and a-galactosidase were
absent. Growth was not significantly inhibited by NaCl concentrations up to 2%, and growth occurred between pH 6.0 and
7.8. The optimum temperature for growth was 37°C, but the
organism also grew at 41°C. No growth was detected at 4°C.
Near-complete (ca. 1,500-bp) small-subunit rRNA genes from
strains 273 and B13 were PCR amplified using eubacteriumspecific primers and conditions previously described (34). Excess amplification primers were removed prior to sequencing
by using commercially available columns per the manufacturer’s instructions (Wizard PCR Preps; Promega, Madison,
Wis.). Double-stranded sequencing of PCR products was performed by automated, fluorescent cycle sequencing, using
modified versions of previously described eubacterium-specific
primers targeted to conserved regions of the 16S rRNA gene
(12, 31, 32, 33). Sequences from the closest relatives of strain
273 were identified and obtained from the Ribosome Database
Project by using the SIMILARITY_RANK and SUBALIGNMENT programs (14), and related sequences in other databases were identified by BLAST analysis. Sequences were then
manually aligned based upon both primary and secondary
structure by using the ARB editor (www.biol.chemie.tumuenchen.de/pub/ARB). The sequences used for phylogenetic
analysis are as follows (GenBank accession numbers given
in parentheses): Pseudomonas sp. strain 273 (AFO30488),
Pseudomonas sp. strain B13 (AFO39489), Pseudomonas aeruginosa LMG1242 (Z76651), Pseudomonas alcaligenes LMG1224
(Z76653), Pseudomonas citronellosis DSM50332 (Z76659),
Pseudomonas flavescens NCPPB3063 (U01916), Pseudomonas fluorescens DSM50090 (Z76662), Pseudomonas mendocina
APPL. ENVIRON. MICROBIOL.
FIG. 1. Phylogenetic tree of Pseudomonas sp. strain 273 and its closest relatives based on 16S RNA sequence analysis.
LMG1223 (Z76664), Pseudomonas nitroreductans IAM1439
(D84006), Pseudomonas oleovorans DSM1045 (Z76665),
Pseudomonas putida DSM291 (Z76667), Pseudomonas stutzeri
CCUG11256 (U25432), Pseudomonas sp. strain IpA-1 (X96787),
and Pseudomonas sp. strain A3 (Y13246). Natural relationships were inferred by neighbor-joining analysis (24). Phylogenetic analyses placed strain 273 in the gamma subgroup of
the Proteobacteria and within the genus Pseudomonas. Its closest relatives were a Pseudomonas species (GenBank accession
no. Y13246) (98.9% similarity), P. citronellolis (98.8% similarity),
and Pseudomonas sp. strain B13 (98.5% similarity). Strain 273
was more distantly related to P. aeruginosa (94.8% similarity)
(Fig. 1). This is consistent with the results of the phenotypic
analyses described above and with a preliminary identification
done by the Deutsche Sammlung von Mikroorganismen und
Zellkulturen (Braunschweig, Germany) based upon the organism’s fatty acid profile and partial rDNA sequence.
1,10-DCD formed a separate phase on top of the aqueous
phase due to its very low solubility in water (,100 mg liter21
[3]). Pseudomonas sp. strain 273 started to grow after 24 h, and
the organic phase disappeared. When baffled Erlenmeyer flasks
were used, 1,10-DCD formed small droplets in the culture medium when shaken at 250 rpm. The same effect was achieved in
regular flasks containing stainless steel spirals (1-cm diameter;
1-mm-thick wire) covering the bottom of the growth vessel.
Alternatively, 1,10-DCD was added to the culture medium as
a 10% emulsion in medium, which had been sonicated until the
suspension was milky. When these techniques were applied,
growth and chloride release started after only 12 h. During
growth of Pseudomonas sp. strain 273 with 1,10-DCD, stoichiometric amounts of chloride were released (Fig. 2). The pH
dropped from 7.4 to 4.7 within 48 h when no pH control was
installed. The cultures began to foam when the substrate was
exhausted, and the pH dropped below 6.2. At pH 4.7, the
culture no longer contained viable cells. Therefore, the pH of
the cultures was routinely adjusted to pH 7.4 with 4 M potassium hydroxide before the pH had dropped below 6.2, which
also prevented foaming. Strain 273 was able to grow on and
release stoichiometric amounts of chloride from a variety of
chlorinated aliphatic compounds, including 1,12-dichlorododecane, 1,10-DCD, 1-chlorodecane, 10-chlorodecane-1-ol, 1,9dichlorononane, 1,8-dichlorooctane, 1,6-dichlorohexane,
and 1,5-dichloropentane. Growth on 1,6-dichlorohexane and 1,5dichloropentane was inhibited when these compounds were
present at concentrations above 2 mM. In contrast, growth and
dechlorination were not influenced by C 8 to C 12 a,vdichloroalkanes, even when present at saturating concentra-
VOL. 64, 1998
AN AEROBIC a,v-DICHLOROALKANE-DEGRADING BACTERIUM
FIG. 2. Batch fermentation of strain 273 in a 10-liter bioreactor with mineral
salts medium containing 5 mM 1,10-DCD as the sole carbon source. The consumption of 1,10-DCD (circles), the release of chloride (triangles), and the
increase in biomass (squares) was monitored over time. pH was kept constant at
7 by automatic titration with NaOH.
tions. Growth was also observed with the following compounds
(concentrations tested given in parentheses): n-heptadecane
(5 mM), n-tetradecane (1 mM), n-dodecane (5 mM), n-decane
(5 mM), phenylnonane (1 mM), n-octane (5 mM), n-hexane
(1 mM), n-pentane (1 mM), 1-eicosanol (1 mM), 1-tetradecanol (1 mM), 1-dodecanol (1 mM), 1-decanol (2 mM), 1octanol (1 mM), 1-hexanol (1 mM), 1,10-decanediol (2 mM),
decanal (5 mM), 2-decanone (5 mM), n-decanoic acid (5 mM),
1,10-decanedioic acid (sebacic acid) (5 mM), 1-decene (2 mM),
n-decanoic acid dimethyl ester (dimethyl sebacate) (2 mM),
10-decanediamide (sebacamid) (5 mM), acetate (10 mM), glucose (10 mM), and glycerol (10 mM). No growth was observed
within 2 weeks with 1 mM isooctane, cyclohexane, cycloheptane, 1,10-diaminodecane, phenol, benzene, toluene, xylene,
and 3-chlorobenzoate. Substances that are solid at ambient
temperatures, like eicosane, did not support growth. In resting
cell experiments with Pseudomonas sp. strain 273 under anaerobic conditions, no chloride release from 1,10-DCD, 1,9-dichlorononane, or 1,8-dichlorooctane was observed, indicating
that oxygen was required for the dechlorinating reactions.
For induction experiments, strain 273 was grown for 48 h in
LC medium (7.15 g of Na2HPO4 z 2H2O, 1.33 g of KH2PO4,
1.0 g of (NH4)2SO4, 0.2 g of MgSO4 z 7H2O, 1 ml of trace element solution, and 1 ml of vitamin solution) containing 10
mM decane or 10 mM 1,10-DCD or in HNB medium. The cells
were harvested by centrifugation at 5,000 3 g for 20 min,
washed once with 25 mM phosphate buffer (pH 7.5), and suspended in 50 ml of either 50 mM phosphate buffer, 50 mM
phosphate buffer containing 100 mg of chloramphenicol ml21
(from an ethanol stock solution of 10 mg ml21), 50 mM phosphate buffer containing 100 mg of kanamycin ml21, or LC medium to give a final concentration of 4 3 108 cells ml21. Each
flask contained 10 mM 1,10-DCD and was incubated at 28°C
and shaken at 200 rpm. Aliquots were taken every hour for to
monitor optical density (A605) and protein and chloride concentrations.
Cells grown on 1,10-DCD immediately dehalogenated 1,10DCD in LC medium with or without chloramphenicol or kanamycin. Cells grown in HNB medium did not dechlorinate 1,10DCD within 12 h, but longer incubation resulted in release of
chloride and growth in the LC medium not amended with
chloramphenicol or kanamycin. No dehalogenation was observed in the medium containing the antibiotics. Control experiments confirmed that the antibiotic concentration was high
enough to inhibit growth. Cells grown with glucose, glycerol, or
acetate dechlorinated 1,10-DCD after a lag phase of 12 h. In
3509
contrast, cells grown on decane dechlorinated 1,10-DCD without a lag phase in the presence and absence of chloramphenicol or kanamycin, suggesting that decane induces the dechlorinating enzyme system. The dehalogenation rates, however, were
lower than those of cells grown on 1,10-DCD. Dechlorination
rates were twice as high in cells grown on 1,10-DCD (25.2 mmol
of chloride released h21 mg of protein21) than in cells grown
on decane (11.3 mmol of chloride released h21 mg of protein21). These results show that the 1,10-DCD-dechlorinating
enzyme system was not constitutive and that the dechlorinating
enzyme system was induced by 1,10-DCD and decane. In the
presence of 5 mM decane and 5 mM 1,10-DCD, the dechlorination rate decreased by 60% (4.5 mmol of chloride h21 mg21
for decane-grown cells and 10.1 mmol of chloride h21 mg21 for
1,10-DCD-grown cells). These results suggest that the enzyme
system responsible for the dehalogenation of 1,10-DCD is the
same enzyme system that initiates the degradation of decane.
These findings were supported by oxygen consumption measurements. Table 1 shows oxygen consumption rates of resting
cells grown with decane or 1,10-DCD. The rate of oxygen
consumption with decane (12.9 mmol of O2 h21 mg21) was
about twofold higher than the rates measured with 1,10-DCD
(5.3 mmol of O2 h21 mg21) in cells grown in decane and 1,10DCD. The oxygen consumption rates are considerably lower
than the rates we determined for chloride release. This discrepancy may be explained by the fact that the mixing of the
insoluble substrates is less efficient in the oxygen measuring
chamber and that the reaction is therefore limited by the transport of the insoluble substrates to the cells. Nevertheless, these
results prove that the enzyme system induced by decane dehalogenates 1,10-DCD and that the enzyme system induced by
1,10-DCD oxidizes decane. The most likely explanation is that
in both cases the same enzyme system is induced. Cells grown
on 10% LB medium did not show any increase of oxygen
consumption with decane or 1,10-DCD but did with 1-decanol.
The presence of chloramphenicol had no effect on oxygen uptake within the 1-h analysis period.
Since Pseudomonas sp. strain 273 grew well on nonchlorinated alkanes, we tested other known alkane degraders for their
ability to grow on 1,10-DCD. The substrate ranges for alkanedegrading organisms differ markedly. Rhodococcus erythropolis
Y2 grew with C7 to C18 n-alkanes and monochloroalkanes, and
the best substrate for dechlorination in resting cell experiments
was 1-chlorotetradecane and 1-chlorohexadecane (1). Scholtz
et al. (25) described an Arthrobacter species which was able to
grow with C4 to C8 a-monochlorinated alkanes, and a hydrolytic dehalogenase was purified from this organism. This enzyme dechlorinated 1-chlorodecane but exhibited no activity
TABLE 1. Oxygen uptake rates of resting cells of Pseudomonas sp.
strain 273 with 1,10-DCD or n-decane as substrates independent of
the presence of protein biosynthesis inhibitors
Growth
substrate
Presence of
antibiotica
1,10-DCD
n-Decane
Oxygen uptake
(mmol h21 mg21)b
1,10-DCD
n-Decane
1
2
5.9
5.9
11.3
12.5
1
2
4.8
4.5
14.2
13.4
a
Cells were incubated in 50 mM phosphate buffer with (1) or without (2) 100
mg of kanamycin or chloramphenicol.
b
The decane- and 1,10-DCD-grown cells used in these experiments had a
basic oxygen demand of 2 to 3 nmol of O2 ml21 min21.
3510
WISCHNAK ET AL.
FIG. 3. Enzymatic conversion of 1,10-DCD (circles) and chloride release
(triangles) in cell extracts of strain 273 in the presence (solid symbols) or absence
(open symbols) of NADH (5 mM) and Fe21 (0.2 mM). Data were averaged from
two independent experiments.
toward 1,10-DCD. Pseudomonas sp. strain C12B grew with C9
to C12 n-alkanes, and optimum growth was observed with C11
n-alkanes, but no dehalogenating activity was reported for this
strain (11). P. oleovorans GPo1 grew with C6 to C12 n-alkanes,
and optimum growth occurred on C8 and C9 n-alkanes (15,
29). Omori and Alexander (19) studied Pseudomonas sp. strain
B which dehalogenated mono- and di-substituted chloroalkanes when grown on n-undecane. In the latter two cases, the
corresponding chlorinated alkanes were cometabolically dehalogenated, but none of them supported growth when supplied
as the only growth substrate. P. oleovorans GPo1 harbors the
OCT plasmid which encodes alkane monooxygenase. Hybridization experiments with the alk genes from this plasmid with
total DNA from Pseudomonas sp. strain 273 revealed no homologous bands. P. oleovorans GPo1 grew well with octane,
decane, 1-chlorodecane, 1-chloro-10-decanol, 1-decanol, and
decanal. However, this organism did not grow with 1,10-DCD.
In the presence of octane as the growth substrate, P. oleovorans
GPo1 released chloride from 1,8-dichlorooctane and 1,9-dichlorononane but not from 1,10-DCD. P. oleovorans DSM
1045T did not grow with any of the a,v-dichloroalkanes. These
results confirm the unique ability of Pseudomonas sp. strain 273
to grow on a,v-chlorinated alkanes, an ability not shared by other known alkane degraders.
Cell extracts of Pseudomonas sp. strain 273 dechlorinated 3
mM 1,10-DCD within 2 h (Fig. 3). The addition of glutathione
(5 mM) had no effect, whereas the divalent metal chelator
EDTA had an inhibitory effect on dechlorination. Only low
dechlorination activities were measured with the soluble protein fraction or with the particulate fraction alone. Complete
dechlorination of the initial amount of 1,10-DCD was achieved
only when both the particulate fraction and the soluble fraction
were added. Furthermore, for the complete dechlorination of
5 mM 1,10-DCD, the addition of NADH and Fe21 was required after dialysis of the cell extract. The reaction was strictly
oxygen dependent, and no dechlorination was observed under
anaerobic conditions. The addition of catalase did not affect
dehalogenation, indicating that a peroxidase was not involved
in the reaction. In the soluble protein fraction obtained from
cells grown on 1,10-DCD, aldehyde dehydrogenase activity was
measured (7) (0.31 U mg of protein21 with decylaldehyde as
the substrate), but no alcohol dehydrogenase activity (8) (with
10-chlorodecane-1-ol as the substrate) could be detected.
Several bacteria are known to contain monooxygenases with
broadsubstratespecificity,e.g.,octanemonooxygenase(28),propane monooxygenase (30), and methane monooxygenase (4).
APPL. ENVIRON. MICROBIOL.
Propane monooxygenase in the presence of oxygen dechlorinated 1-chlorobutane, but 1-chlorobutane was not an inducer for either enzyme. This may be the reason why it is
difficult to find dechlorinating activities in microcosms when
chloroalkanes are used as the only substrate. For example, P.
aeruginosa S7B1, isolated and grown with n-hexadecane, aerobically dechlorinated 1,10-DCD (20). However, attempts by
these researchers to culture for dechlorinating organisms with
chloroalkanes as the only growth substrate failed. These findings suggest that enzymes involved in the initial catabolic step
of n-alkane degradation also play a role in chloroalkane dechlorination. However, these enzymes were induced only by
the nonchlorinated alkane and not by the corresponding chloroalkane. Similar data were reported for an oxygenase-type dehalogenase from R. erythropolis Y2 (1). Hexadecane-grown resting cells of R. erythropolis Y2 did dechlorinate 1,10-DCD, but
no growth occurred on the a,v-chlorinated compound. In contrast, dechlorinating activity in Pseudomonas sp. strain 273 was
induced by both 1,10-DCD and decane. The v-hydroxylating
multienzyme system involved in the hydroxylation of n-alkanes has been characterized from octane-grown cells of P.
oleovorans (16, 22, 23). This enzyme system is composed of
rubredoxin, NADH-rubredoxin reductase, and the nonheme
ion v-hydroxylase (a monooxygenase). v-Hydroxylase has
been described as a relatively insoluble and unstable protein
that required the presence of ferrous ion and phosholipids for
full activity (22, 23). At this stage, it seems likely that the
a,v-chloroalkane-dechlorinating enzyme system from strain
273 is a monooxygenase with a composition similar to that of
the well-known v-hydroxylase.
Nucleotide sequence accession numbers. The 16S ribosomal
DNA (rDNA) sequences of Pseudomonas sp. strain 273 and
Pseudomonas sp. strain B13 were deposited as GenBank accession numbers AFO39488 and AFO39489, respectively.
This research was supported by a Feodor-Lynen fellowship from the
Alexander von Humboldt-Stiftung to F.E.L. and by National Science
Foundation grant DEB9120006 to the Center for Microbial Ecology.
The technical assistance by Waltraut Rüde and Christine Lach during the initial isolation of Pseudomonas sp. strain 273 is gratefully acknowledged. We also thank J. van Beilen and M. Schlömann for providing Pseudomonas oleovorans GPol and Pseudomonas sp. strain B13,
respectively; G. Auling and U. Griepenburg for performing the polyamine analyses; and Jan Rademaker for his help with the BOX- and
ERIC-PCR analyses. We are indebted to Jim Tiedje for his support
and many helpful suggestions.
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