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Applied and Environmental Microbiology, December 1999, p. 5607-5611, Vol. 65, No. 12
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Isolation of Terrabacter sp. Strain
DDE-1, Which Metabolizes 1,1-Dichloro-2,2-Bis(4-Chlorophenyl)Ethylene
when Induced with Biphenyl
J.
Aislabie,1,*
A. D.
Davison,2,
H. L.
Boul,3
P. D.
Franzmann,4
D. R.
Jardine,5 and
P.
Karuso5
Landcare Research,
Hamilton,1 and Wool Research
Organisation of New Zealand Inc., Christchurch,3
New Zealand, and Key Centre for Biodiversity and
Bioresources2 and School of
Chemistry,5 Macquarie University, Sydney
2109, and CSIRO Land and Water, Floreat Park, WA
6014,4 Australia
Received 10 May 1999/Accepted 14 September 1999
 |
ABSTRACT |
Terrabacter sp. strain DDE-1, able to metabolize
1,1-dichloro-2,2-bis(4-chlorophenyl)ethylene (DDE) in pure culture when
induced with biphenyl, was enriched from a
1-1-1-trichloro-2,2-bis(4-chlorophenyl)ethane residue-contaminated
agricultural soil. Gas chromatography-mass spectrometry analysis of
culture extracts revealed a number of DDE catabolites, including
2-(4'-chlorophenyl)-3,3-dichloropropenoic acid,
2-(4'-chlorophenyl)-2-hydroxy acetic acid, 2-(4'-chlorophenyl) acetic
acid, and 4-chlorobenzoic acid.
| |
TEXT |
The insecticide
1,1,1-trichloro-2,2-bis(4-chlorophenyl)ethane, commonly known
as DDT, was once used extensively to control both agricultural pests
and disease vectors. Although many countries ceased using DDT over 20 years ago, its residues (DDTr) still persist in the environment,
predominantly as 1,1-dichloro-2,2-bis(4-chlorophenyl)ethylene (DDE), 1,1-dichloro-2,2-bis(4-chlorophenyl)ethane
(DDD), and
1,1,1-trichloro-2-(2-chlorophenyl)-2-(4-chlorophenyl)ethane (2,4'-DDT), as well as unchanged DDT (Fig.
1) (1). Among these residues,
DDE is the most toxicologically significant residue, as it blocks the
action of androgens in rats (12) and has been implicated in
male reproductive abnormalities in alligators (9). In
topsoils, the presence of DDTr, often as DDE, limits land use options
and may have an impact on trade in agricultural products. Cost-effective remediation methods for widespread low-level
contamination are therefore required. While bacterial degradation of
DDE has recently been reported in a recombinant strain (10),
it is not known how widespread this capacity is in nature. The presence of organisms with this degradative capacity in contaminated soil may be
beneficial in developing treatment options.
DDE has been considered to be a dead-end metabolite of DDT formed under
oxidizing conditions (1, 8). Although little is known about
the microbial metabolism of DDE (1, 8, 16), recent reports
indicate that it can be metabolized by chlorobiphenyl-degrading bacteria under aerobic conditions (10, 13). Bacterium strain B-206, for example, produced a number of phenolic metabolites from DDE
(13), and subsequently, Pseudomonas acidovorans
M3GY was shown to mediate ring cleavage of DDE proceeding via
meta-fission to 4-chlorobenzoic acid (10).
To investigate whether aerobic bacteria from DDTr-contaminated
agricultural soils have the potential to metabolize DDE, we established
enrichment cultures to select for microbes that transform DDE when
provided with biphenyl as a carbon source. An aerobic gram-positive
bacterium, Terrabacter sp. strain DDE-1, which metabolizes DDE when grown with biphenyl, was isolated.
Chemicals.
DDE was purchased from Aldrich Chemical Company,
2,4'-DDT was purchased from Riedel de Haen, and biphenyl was purchased
from BDH. [U-14C]DDE, prepared from
[U-14C]DDT, was obtained from NEN Research Products,
Boston, Mass. (4). All chemicals were at least 98% pure.
Diazomethane was generated from
N-methyl-N-nitroso-p-toluenesulfonamide
with the Diazald kit (Aldrich Chemical Company) according to the
manufacturer's instructions.
Enrichment cultures.
Enrichment cultures for the selection of
microbes metabolizing DDE were established in 100 ml of minimal medium
(18) with the addition of vitamins (17) and
biphenyl (1 mg ml
1) and in the presence of DDE (0.25 mg
ml1). Biphenyl and DDE dissolved in a minimal amount of
acetone were added to sterile flasks, the solvent was evaporated, and
then sterile medium was added. The source of inocula (10 g per flask) was agricultural soil from the AgResearch Winchmore Research Station, Canterbury, New Zealand, in which levels of DDE may reach 2 mg kg1. Details of these soils are described by Boul et al.
(3). All enrichment cultures were incubated at 28°C on a
rotary shaker at 200 rpm. Aliquots (10 ml) were subcultured in fresh
medium every month. Growth was indicated by the increase in turbidity in enrichment cultures with biphenyl as a carbon source in the presence
of DDE. After 3 months, a persistent yellow water-soluble product
accumulated in one of the cultures. Removal of DDE from the culture
accumulating the yellow product was revealed by high-pressure liquid
chromatography analysis of solvent extracts from entire cultures after
1 month of incubation (4). There was no loss of DDE or
accumulation of a yellow product in the abiotic control flasks.
Isolation and identification of a DDE-metabolizing bacterium.
A bacterium which metabolizes DDE was isolated from the enrichment
culture by an overlayer technique (19). The DDE precipitated in the overlayer medium formed an opaque layer, and biphenyl crystals were placed in the lid of the petri dishes. Presumptive
DDE-metabolizing microbes were detected as colonies surrounded by a
clear zone after at least 6 weeks of incubation at 28°C. Isolates
were removed from the clear zones and purified by streaking them onto
plate count agar (PCA) (Difco). A bacterium, designated strain
DDE-1, was selected for further study. Within 48 h of incubation,
it produced a yellow water-soluble product on PCA which had previously been spread with 0.1 ml of 1% (wt/vol) DDE in ether.
Strain DDE-1 is a gram-positive, coccobacillus-shaped bacterium which
is catalase positive, oxidase negative, and nonmotile.
16S rRNA
sequence analysis (
2,
6) showed that strain DDE-1
belonged
within
Terrabacter, a genus of the high-G+C gram-positive
bacterial clade. Strain DDE-1 was 98.6% similar to
Terrabacter sp. strain DPO1361 and 95.9% similar to
Terrabacter tumescens.
A phylogenetic tree was constructed
(
7,
20), and the position
of strain DDE-1 is shown in Fig.
2.
Terrabacter sp. strain
DDE-1
has been deposited in the International Collection of
Micro-organisms
from Plants (ICMP; Auckland, New Zealand) as strain
ICMP 13121.
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FIG. 2.
16S rRNA phylogenetic tree showing the position of
Terrabacter sp. strain DDE-1. The GenBank accession numbers
of the organisms used were as follows: Terrabacter sp.
strain DPO1361, Y08853; T. tumescens NCIB 8914, X53215;
Dermatophilus congolensis ATCC 14637, M59057;
Dermabacter hominis DSM 7083, X91034; Clavibacter
xyli clone pCG803, M60935; Clavibacter michiganensis
DSM 7483, X77434; Brachybacterium faecium DSM 4810, X91032;
Brachybacterium alimentarium CNR2 925, X91031;
Kytococcus sedentarius DSM 20547, X87755; Kytococcus
nishinomiyaensis DSM 20448, X87757; and Nocardia
asteroides ATCC 19247, Z36934. The bar represents a Jukes-Cantor
distance of 0.05.
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Evidence for the metabolism of DDE by Terrabacter sp.
strain DDE-1.
Metabolism of DDE by Terrabacter sp.
strain DDE-1 in pure culture was confirmed by the loss of DDE from
culture flasks containing minimal medium and biphenyl. Cells grown on
PCA with biphenyl in the lid were removed from the agar, washed twice,
and resuspended in phosphate buffer, and 2.5 × 109
cells were inoculated into a series of flasks containing 50 ml of
minimal medium supplied with biphenyl (0.5 mg ml1) and
DDE (0.1 mg ml1). Biotic control flasks were inoculated
with cells killed by autoclaving. All cultures were incubated at 28°C
in the dark with shaking at 200 rpm. Entire flasks were sacrificed at
regular intervals. The numbers of viable cells were monitored by
plating culture dilutions on PCA. Triplicate cultures were acidified
with 0.5 ml of concentrated HCl, 2,4' DDT (5 mg per flask) was added as an internal standard, and DDE residues were extracted with hexane. The
extracts were diluted 100-fold in hexane and analyzed with an Econo-Cap
EC-1 nonpolar silica column coated with polydimethylsiloxane (30 m by
0.32 mm [internal diameter]; 0.25-µm film thickness; Alltech) and a
Shimadzu model GC17AV2 gas chromatograph with a wide-range flame
ionization detector. The operating conditions were as follows: injector
port, 250°C; detector, 270°C; column temperature program,
50°C isothermal for 1 min, 10°C/min to 200°C, 20°C/min to
300°C, and isothermal at 300°C for 5 min. The carrier gas was
helium at 1 ml/min with a pressure program, and injections were 1 µl splitless.
DDE was lost from the flasks containing live
Terrabacter sp.
strain DDE-1 but not from the abiotic control flasks after 28
days of
incubation (Fig.
3). The concentration of
DDE fell from
0.1 to 0.062 mg ml
1 after 10 days of
incubation, after which no further loss occurred.
While no metabolites
of DDE were detected in the extracts by using
a wide-range flame
ionization detector, the accumulation of a
yellow product in the
medium, indicative of ring cleavage, was
observed, as noted previously.
No concomitant increase in viable
cell numbers was detected by
plating culture dilutions onto PCA
(results not shown).
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FIG. 3.
Rate of transformation of DDE by Terrabacter
sp. strain DDE-1 when incubated with biphenyl ( ). Control flasks
containing dead cells are also shown ( ). The data are the means of
values from three replicates; the bars indicate standard errors.
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Additional evidence for the metabolism of DDE by
Terrabacter
sp. strain DDE-1 was obtained with [
14C]DDE. Biometer
flasks containing 30 ml of minimal medium supplemented
with DDE (0.1 mg
ml
1) spiked with 0.225 µCi of [phenyl
ring-
14C(U)]DDE were inoculated with cells induced with
biphenyl as described
above. Potassium hydroxide (1 M) was used as a
CO
2 trap, and the
flasks were incubated at room temperature
in the dark with shaking
at 125 rpm. The experiment was carried out in
triplicate with
biotic and abiotic controls. At regular intervals,
0.5-ml samples
of the CO
2 trap were removed and mixed with
11 ml of the scintillation
cocktail (10 ml of PCS plus 1 ml of water),
and the radioactivity
was determined by liquid scintillation counting
with a Beckman
model 3800 liquid scintillation counter. The
14C residues in the neutral and acid organic phases, the
cell biomass,
and the aqueous phase of the cultures were quantified as
follows.
The cultures were adjusted to pH 7.5 with 5 M NaOH and
extracted
three times with an equal volume of ethyl acetate. The
remaining
culture was adjusted to pH 2.5 to 3.5 with 5 M HCl and
reextracted
with ethyl acetate as before. The culture was then
centrifuged,
and the pellet of cells was washed and then resuspended in
10
ml of buffer, from which a 1-ml aliquot was removed for
scintillation
counting. The radioactivity present in 1-ml aliquots of
the supernatant
combined with cell washings and in 1-ml aliquots of the
solvent
extracts was also
measured.
A mass balance calculated for
Terrabacter sp. strain DDE-1
(Table
1) reveals conclusively the
ability of this bacterium to
metabolize DDE. In comparison with the
control flasks, those containing
live cells transformed a significant
percentage of the [
14C]DDE to acid-extractable products.
At the same time, the percentage
of radioactivity recoverable from the
neutral extract decreased
in flasks with live cells. No mineralization
of DDE to carbon
dioxide was detected. Similar levels of conversion of
DDE and
the absence of mineralization activity were reported for
P. acidovorans M3GY (
10).
The spectral characteristics of the yellow product were obtained by
inoculating washed cells of
Terrabacter sp. strain DDE-1,
induced with biphenyl as above, into minimal medium containing
DDE (0.1 mg ml
1) only. The flasks were incubated at room
temperature in the dark
with shaking at 125 rpm for 10 days. An aliquot
of 3 ml was taken
from the flask and centrifuged, and the spectrum of
the supernatant
was determined over the range from 350 to 550 nm in a
Beckman
DU-640 spectrophotometer. The yellow compound had an absorbance
maximum at 401 nm at neutral pH; acidification with orthophosphoric
acid caused a shift in the spectrum and a concomitant decrease
in the
peak. These features are characteristic of
meta-ring
cleavage
products of aromatic compounds that absorb over the 375 to
430-nm
region (
10,
14). A proposed structure for this
product (catabolite
II) is given in Fig.
4.
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FIG. 4.
Catabolites identified in the degradation of DDE by
Terrabacter sp. strain DDE-1 by GC-MS (methyl ester
derivatives) and UV-visible-light spectrophotometry. ND, not visualized
by GC-MS; +, present.
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Isolation and characterization of DDE catabolites.
A number of
DDE catabolites were isolated from cultures of Terrabacter
sp. strain DDE-1 in minimal medium with DDE (0.5 mg ml1)
and induced with biphenyl (Fig. 4). To this end, the cultures were
incubated in the dark at 25°C with shaking at 180 rpm for 7 or 21 days, acidified to pH 2 to 3 with orthophosphoric acid, and centrifuged
(10,000 × g; 30 min; 4°C) to remove cells and debris. The supernatant was extracted with approximately 1.5 volumes of
ethyl acetate. The extract was dried over anhydrous sodium sulfate and
concentrated to approximately 500 µl in vacuo with gentle heat
(30°C). The concentrated sample was further reduced under a stream of
nitrogen and methylated with diazomethane before gas
chromatography-mass spectrometry (GC-MS) analysis (models GC 8000 and
MD 800; Fison's Instrument). The gas chromatograph was fitted with a
BPX5 column coated with polysilphenylene-siloxane (length, 25 m;
internal diameter, 0.22 mm; 0.25-µm film thickness). Injection (2 µl) was performed in split mode at 40°C. The column temperature was
held at 40°C for 2 min and then raised by 5°C per min up to
280°C, where it was held for 5 min. The injector temperature was
maintained at 250°C. The carrier gas was helium at a pressure of 80 kPa.
Mass spectra were obtained (electron ionization at 70 eV; trap current,
100 µA; mass range, 40 to 500; source temperature,
200°C), and
possible structures were deduced by comparing the
mass spectra recorded
with the mass spectrum database of the National
Institute of Standards
and Technology and reports in the literature
(
10,
14).
GC-MS analyses of the culture extracts revealed many catabolites (Fig.
4) that were not detected in abiotic controls. The
presence of chlorine
in the catabolites was confirmed by analysis
of the molecular ion.
Compounds containing one chlorine atom typically
show a molecular ion
with an
m/z of
x + 2 in a ratio of 3 to 1
that corresponds to the two naturally occurring chlorine isotopes
(35 and 37). This feature helps confirm DDE as the parent compound
from
which the catabolite is derived (
5).
Two compounds with different retention times (44.5 and 45.3 min,
respectively) but identical mass spectra and molecular ions
with
m/z of 332 (16 mass units more than the parent molecule)
were detected (Fig.
5). The major
fragments of the ionization
pattern were indicative of the loss of one
chlorine (
m/z, 297),
two chlorines (
m/z, 262),
and HCl (
m/z, 296) (Fig.
5). These catabolites
were
identified as isomers of the monohydroxylated derivatives
(catabolite I
[Fig.
4]) of DDE at the
meta and
ortho
positions,
based on comparison with the database reference spectra and
similarity
to the spectrum presented by Hay and Focht (
10).
We propose
that the isomers are derived from dehydration of an initial
dihydroxy
compound, formed through dioxygenase attack on DDE, which is
unstable
in acidic extracts (
10,
14).
A number of catabolites downstream of the proposed ring cleavage
product (catabolite II) were isolated. Catabolite III (Fig.
6), identified as
2-(4'-chlorophenyl)-3,3-dichloropropenoic acid
(methyl ester), had a
molecular ion at an
m/z of 264. This compound
was also
reported as a metabolite of DDE when degraded by
P. acidovorans M3GY (
10). No catabolites that were
intermediate between compounds
II and III were seen in the GC-MS
analysis. Catabolite IV, 2-(4'-chlorophenyl)-2-hydroxy
acetic acid
(methyl ester), with a molecular ion with an
m/z of
200, represents a loss of carbon dioxide and HCl from catabolite
III,
indicating sequential degradation. This compound could then
be further
degraded to 4-chlorobenzoic acid (catabolite VI [methyl
ester]).
Catabolite V (
m/z, 184; methyl ester) is also likely
to be
formed from catabolite IV, possibly through dehydration,
but may be a
dead-end product in this pathway. Catabolites V and
VI have also
been reported previously as degradation products
of DDT (
14,
15) and DDE (
10).
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FIG. 6.
Mass spectrum for catabolite III,
2-(4'-chlorophenyl)-3,3-dichloropropenoic acid (methyl ester).
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In conclusion, a gram-positive bacterium,
Terrabacter sp.
strain DDE-1, was isolated that is able to metabolize DDE to
4-chlorobenzoate
when induced with biphenyl. The evidence points to the
involvement
of a dioxygenase and subsequent
meta-ring
cleavage similar to
the degradation pathway proposed for
P. acidovorans M3GY (
10).
This is only the second report
of extensive metabolism of DDE
and the first report of this metabolism
in a gram-positive organism.
As
Terrabacter sp. strain DDE-1
was isolated from an agricultural
soil contaminated with DDE, this
study provides evidence for the
usefulness of biphenyl-metabolizing
bacteria for in situ remediation
of DDE-contaminated agricultural
soils. To enhance degradation
of DDE in soils, information is required
on the in situ abundance
and distribution of bacteria with this type of
metabolism and
the substrates that will induce the expression of the
catabolic
pathway for DDE degradation. While biphenyl has proved a
useful
substrate for isolating
Terrabacter sp. strain DDE-1,
the application
of biphenyl to soils is not desirable, as it is on the
Environmental
Protection Agency's priority pollutant list.
Fortunately, a number
of plant terpenes, including cymene and limonene,
have been shown
to stimulate degradation of polychlorinated biphenyls
(
11).
Whether these compounds also induce DDE degradation in
Terrabacter sp. DDE-1 has yet to be
determined.
Although remediation of DDTr-contaminated soils is difficult, the
isolation of a bacterium with the ability to degrade DDE
from such a
site indicates that microbially mediated processes
for cleanup of
DDTr-contaminated soils are worthy of further
investigation.
Nucleotide sequence accession number.
The 16S rRNA sequence of
strain DDE-1 has been submitted to GenBank and assigned accession no.
U96645.
| |
ACKNOWLEDGMENTS |
This study was supported by the Foundation for Research, Science
and Technology (New Zealand) under contract C10 546 and by a NSW
Environmental Trust Research Grant (Australia). A.D.D. is funded by a
Macquarie University Research Fellowship.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Landcare
Research, Private Bag 3127, Hamilton, New Zealand. Phone: 64 7 858-3700. Fax: 64 7 858-4964. E-mail:
aislabiej{at}landcare.cri.nz.
Present address: Australian Water Technologies, W. Ryde, NSW 2114, Australia.
| |
REFERENCES |
| 1.
|
Aislabie, J. M.,
N. K. Richards, and H. L. Boul.
1997.
Microbial degradation of DDT and its residues a review.
N. Z. J. Agric. Res.
40:269-282.
|
| 2.
|
Blackall, L. L.
1994.
Molecular identification of activated sludge foaming bacteria.
Water Sci. Technol.
29:35-42.
|
| 3.
|
Boul, H. L.,
M. L. Garnham,
D. Hucker,
D. Baird, and J. Aislabie.
1994.
Influence of agricultural practices on levels of DDT and its residues in soil.
Environ. Sci. Technol.
28:1397-1402.
|
| 4.
|
Boul, H. L.
1996.
Effect of soil moisture on the fate of radiolabelled DDT and DDE in vitro.
Chemosphere
32:855-866.
|
| 5.
|
Davison, A. D.,
P. Karuso,
D. R. Jardine, and D. A. Veal.
1996.
Halopicolinic acids, novel products arising through the degradation of chloro- and bromobiphenyl by Sphingomonas paucimobilis BPS1-3.
Can. J. Microbiol.
42:66-71[Medline].
|
| 6.
|
Dorsch, M., and E. Stackebrandt.
1992.
Some modifications in the procedure of direct sequencing of PCR amplified 16S rDNA.
J. Microbiol. Methods
16:271-279.
|
| 7.
|
Felsenstein, J.
1989.
PHYLIP-phylogeny inference package (version 3.2).
Cladistics
5:164-166.
|
| 8.
|
Focht, D. D.
1993.
Microbial degradation of chlorinated biphenyls, p. 341-407.
In
J.-M. Bollag, and G. Stotzky (ed.), Soil biochemistry, vol. 8. Marcel Dekker, Inc., New York, N.Y
|
| 9.
|
Guillette, L. J.,
D. B. Pickford,
D. A. Crain,
A. A. Rooney, and H. F. Percival.
1996.
Reduction in penis size and plasma testosterone concentrations in juvenile alligators living in a contaminated environment.
Gen. Comp. Endocrinol.
101:32-42[Medline].
|
| 10.
|
Hay, A. G., and D. D. Focht.
1998.
Cometabolism of 1,1-dichloro-2,2-bis(4-chlorophenyl)ethylene (DDE) by Pseudomonas acidovorans M3GY grown on biphenyl.
Appl. Environ. Microbiol.
64:2141-2146[Abstract/Free Full Text].
|
| 11.
|
Hernandez, B. S.,
S. C. Koh,
M. Chial, and D. D. Focht.
1997.
Terpene-utilizing isolates and their relevance to enhanced biotransformation of polychlorinated biphenyls in soil.
Biodegradation
8:153-158.
|
| 12.
|
Kelce, W. R.,
C. R. Stone,
S. C. Laws,
L. E. Gray,
J. A. Kemppainen, and E. M. Wilson.
1995.
Persistent DDT metabolite p,p'-DDE is a potent androgen receptor antagonist.
Science
375:581-585.
|
| 13.
|
Massé, R.,
D. Lalanne,
F. Meisser, and M. Sylvestre.
1989.
Characterization of new bacterial transformation products of 1,1,1-trichloro-2,2-bis-(4-chlorophenyl)ethane (DDT) by gas chromatography/mass spectrometry.
Biomed. Environ. Mass. Spectrom.
18:741-752[Medline].
|
| 14.
|
Nadeau, L. J.,
F.-M. Menn,
A. Breen, and G. S. Sayler.
1994.
Aerobic degradation of 1,1,1-trichloro-2,2-bis(4-chlorophenyl)ethane (DDT) by Alcaligenes eutrophus A5.
Appl. Environ. Microbiol.
60:51-55[Abstract/Free Full Text].
|
| 15.
|
Pfaender, F. K., and M. Alexander.
1972.
Extensive microbial degradation of DDT in vitro and DDT metabolism by natural communities.
J. Agric. Food Chem.
20:842-846[Medline].
|
| 16.
|
Quensen, J. F., III,
S. A. Mueller,
M. K. Jain, and J. M. Tiedje.
1998.
Reductive dechlorination of DDE to DDMU in marine sediment microcosms.
Science
280:722-724[Abstract/Free Full Text].
|
| 17.
|
Roger, P.,
A. Erben, and F. Lingens.
1990.
Microbial metabolism of quinoline and related compounds. IV. Degradation of isoquinoline by Alcaligenes faecalis Pa and Pseudomonas diminuta.
Biol. Chem. Hoppe-Seyler.
371:511-513[Medline].
|
| 18.
|
Rosenberger, R. F., and S. R. Elsden.
1960.
The yields of Streptococcus faecalis grown in continuous culture.
J. Gen. Microbiol.
22:726-739.
|
| 19.
|
Shotbolt-Brown, J.,
D. W. F. Hunter, and J. Aislabie.
1996.
Isolation and description of carbazole-degrading bacteria.
Can. J. Microbiol.
42:79-82[Medline].
|
| 20.
|
Swofford, D. L.
1990.
PAUP: phylogenetic analysis using parsimony, version 3.1.
Illinois Natural History Survey, Champaign, Ill
|
Applied and Environmental Microbiology, December 1999, p. 5607-5611, Vol. 65, No. 12
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
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