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Appl Environ Microbiol, January 1998, p. 153-158, Vol. 64, No. 1
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Morpholine Degradation Pathway of Mycobacterium aurum
MO1: Direct Evidence of Intermediates by In Situ 1H
Nuclear Magnetic Resonance
B.
Combourieu,1
P.
Besse,1
M.
Sancelme,1
H.
Veschambre,1
A. M.
Delort,1,*
P.
Poupin,2 and
N.
Truffaut2
Laboratoire de Synthèse,
Electrosynthèse et Etude de Systèmes à
Intérêt Biologique, UMR 6504 CNRS, Université
Blaise Pascal, 63177 Aubière Cedex,1 and
Laboratoire de Génétique Microbienne,
Université de Technologie de Compiègne, B.P. 649, 60206 Compiègne,2 France
Received 23 June 1997/Accepted 31 August 1997
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ABSTRACT |
Resting Mycobacterium aurum MO1 cells were incubated
with morpholine, a waste from the chemical industry. The kinetics of biodegradation was monitored by using in situ nuclear magnetic resonance (NMR). The incubation medium was directly analyzed by 1H NMR. This technique allowed the unambiguous
identification of two intermediates of the metabolic pathway involved
in the biodegradation process, glycolate and 2-(2-aminoethoxy)acetate.
The latter compound, which was not commercially available, was
synthesized, in three steps, from 2-(2-aminoethoxy)ethanol.
Quantitative analysis of the kinetics of degradation of morpholine was
performed by integrating the signals of the different metabolites in
1H-NMR spectra. Morpholine was degraded within 10 h.
The intermediates increased during the first 10 h and finally
disappeared after 20 h incubation. Assays of degradation were also
carried out with glycolate and ethanolamine, hypothetical intermediates
of the morpholine degradation pathway. They were degraded within 4 and 8 h, respectively. Until now, no tool for direct detection of intermediates or even morpholine has been available, consequently, only
hypothetical pathways have been proposed. The approach described here
gives both qualitative and quantitative information about the metabolic
routes used in morpholine degradation by M. aurum MO1. It
could be used to investigate many biodegradative processes.
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INTRODUCTION |
Recent studies have shown the
biodegradative abilities of Mycobacterium species: they are
able to metabolize xenobiotics such as polycyclic aromatic hydrocarbons
(4, 10, 13, 14, 29), chlorophenols (11),
trichloroethylene (33), vinyl chloride (12),
amines (6), and isonicotinate (18).
Mycobacterium spp. were also found to attack the
heterocyclic, secondary amine morpholine
(C4H9NO), which had been considered to be
persistent for many years. This chemical has great industrial
importance and a wide range of applications (22); it is used
in the manufacture of rubber additives and also as a very versatile
solvent, as an anticorrosive agent, and in the production of various
drugs and pesticides. Its high solubility in water and its high
potential for N nitrosation, which gives the potent mutagen and
carcinogen N-nitrosomorpholine (9, 21, 27), make
this xenobiotic of special interest from an environmental point of
view. Knapp et al. (15) first discovered two strains of
Mycobacterium (MorD and MorG) that were able to utilize
morpholine as a sole source of carbon, nitrogen, and energy. A few
years later, Dmitrenko et al. (5) isolated a strain of
Arthrobacter, and Cech et al. (3) found a
strain of Mycobacterium aurum MO1 that had morpholine degradation properties. Knapp's group studied other
Mycobacterium strains isolated from activated sludges
(1, 16, 17). In the companion paper (24), we
describe another Mycobacterium strain that is able to
degrade morpholine.
More recently, a few studies were carried out in order to understand
the morpholine biodegradation process and its regulation. Swain et al.
(28) proposed a hypothetical pathway for the biodegradation of morpholine by Mycobacterium chelonae (Fig.
1) that could proceed via
2-(2-aminoethoxy)acetate and glycolate and/or ethanolamine. Mazure and
Truffaut (19) described the degradation of morpholine by
M. aurum MO1. They proposed that M. aurum grown
on morpholine could degrade intermediary compounds via the ethanolamine
and glycolate routes. Depending on the morpholine concentration in the
medium, one pathway could be used while the other was inhibited.
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FIG. 1.
Hypothetical morpholine biodegradation pathway of
M. chelonae proposed by Swain et al. (28). CoA,
coenzyme A.
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However, to date, no tool for the direct detection of intermediates, or
even of morpholine, has been available. Only indirect strategies were
developed, such as chemical oxygen demand, optical density, or
NH3 measurements, growth on intermediates, or in vitro enzyme assays. Consequently, only hypothetical pathways were proposed, and limited interpretations of various experiments were made.
In this work we describe the degradation of morpholine by M. aurum MO1 by using in situ 1H nuclear magnetic
resonance (1H-NMR) spectroscopy. This technique has been
previously applied in medical fields for analyzing biological fluids
(20, 23) and also for studying microbial physiology with
extracts or culture medium (8, 30, 31). More recently,
Gaines et al. (7) described the first use of
1H-NMR spectroscopy to monitor the catabolism of mixtures
of aromatic compounds by Acinetobacter calcoaceticus grown
in fully deuterated medium. This method is very interesting and allowed
the identification and quantification of metabolites which
accumulated during growth by rendering invisible the fully
deuterated microbial cultures. However, it cannot be used in all
circumstances, because it requires specialized growth medium and large
quantities of D2O. In this work, we studied the degradative
metabolism of M. aurum MO1 by directly analyzing the
incubation medium in H2O. Our method, using normal water,
can be more generally used and does not perturb the system being
studied (the deuterated compounds can affect the enzymatic reactions).
1H-NMR spectroscopy is both qualitative and quantitative,
so it allowed us to establish unambiguously some steps of morpholine degradation by this strain.
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MATERIALS AND METHODS |
Chemicals.
Morpholine, sodium acetate,
2-(2-aminoethoxy)ethanol, and glycolic acid were purchased from Aldrich
Chemical (Sigma Aldrich Sarl, St. Quentin Fallavier, France),
ethanolamine was purchased from Prolabo (Vault en Velin, France), and
tetradeuterated sodium trimethylsilylpropionate (TSPd4) was
purchased from EurisoTop (St. Aubin, France).
Growth conditions.
M. aurum MO1 cultures were
grown in 100 ml of Trypticase soy broth (bioMérieux,
Marcy-L'Etoile, France) in 500-ml Erlenmeyer flasks incubated at
30°C with agitation at 200 rpm. They were harvested after 48 h
of culture.
Incubation with xenobiotics.
Cells were harvested by
centrifugation at 9,000 × g for 15 min at 5°C. The
supernatant was eliminated, and the pellet was washed twice with Knapp
buffer (containing, per liter of distilled water,
KH2PO4 [1 g], K2HPO4
[1 g], FeCl3 [4 mg], and MgSO4 · 7H2O [40 mg], pH 6.6) and finally resuspended in this
buffer (5 g of wet cells in 50 ml of buffer). The cells were incubated
with 10 mM morpholine, 10 mM ethanolamine, or 13 mM glycolic acid as
the only source of energy in a 500-ml Erlenmeyer flask at 30°C with agitation (200 rpm). Incubation of cells under the same conditions in
the absence of substrate constituted a negative control, as did
incubation of the substrate in the buffer without cells. Samples (1 ml)
were taken every hour for 12 h and from time to time until 72 h. They were centrifuged at 12,000 × g for 5 min. The
supernatants were isolated and immediately frozen until NMR analysis.
1H-NMR spectroscopy. (i) Preparation of NMR
samples.
The supernatant (540 µl) was supplemented with 60 µl
of a 8 mM solution of TSPd4 in D2O and adjusted
to pH 10 with 4 N NaOH. pH adjustment avoided changes in chemical
shifts. D2O was used for locking and shimming.
TSPd4 constituted a reference for chemical shifts (0 ppm)
and quantification.
(ii) 1H-NMR spectra.
1H NMR was
performed at 300.13 MHz on a 300 MSL Bruker spectrometer at 21°C with
5-mm-diameter tubes containing 500 µl of sample; water was suppressed
by saturation with a classical NOE Bruker program. About 150 scans were
collected (90° pulse, 3.7 µs; relaxation delay, 6 s;
acquisition time, 1.024 s; 8,000 data points; saturation time, 2 s). No filter was applied before Fourier transformation, but a baseline
correction was performed on spectra before integration with Bruker
software. Under these conditions, the limit of quantification was in
the range of 0.05 mM.
(iii) Quantification of metabolites.
The concentration of
metabolites was calculated as follows: [m] = (9A0 × [TSPd4])/(b × Aref), where [m] is the concentration of
metabolite m, A0 is the area of metabolite m
resonance in the 1H-NMR spectrum, [TSPd4] is
the concentration of the reference, Aref is the
area of reference resonance in the 1H-NMR spectrum,
b is the number of protons of metabolite m in the signal
integrated, and 9 is the number of protons resonating at 0 ppm.
Synthesis of 2-(2-aminoethoxy)acetate. (i) Protection of the
amino function.
The reaction was conducted as described previously
(25). To a solution of 500 mg (4.8 mmol) of
2-(2-aminoethoxy)ethanol in 10 ml of ethyl acetate, stirred at room
temperature, was added 1.25 g (1.2 equivalents, 5.8 mmol) of
di-tert-butyldicarbonate. The mixture was stirred overnight
at room temperature. The solvent was then evaporated under vacuum, and
the residue was purified by column chromatography (the eluent was
pentane-ether [50:50, vol/vol]). The yield was 92%, with
1H NMR (400.13 MHz, CDCl3) 
ppm 1.40 (s,
9H), 3.15 (s, 1H), 3.25 (t, 2H, J = 5 Hz), 3.46 to 3.56 (m, 4H), 3.75 (t, 2H, J = 4 Hz), 5.30 (s, 1H); 13C NMR
(100.62 MHz, CDCl3) ppm 27.9 (C-7), 39.8 (C-4), 60.7 (C-1), 69.7 (C-3), 71.8 (C-2), 78.4 (C-6), 155.9 (C-5).
(ii) Oxidation of the N-protected alcohol.
The oxidation of
the N-protected alcohol was performed as previously described
(2). A flask was charged with 4.9 ml of carbon
tetrachloride, 4.9 ml of acetonitrile, 7.3 ml of water, 0.5 g
(2.45 mmol) of the N-protected alcohol, and 2.14 g (4.1 equivalents, 10 mmol) of sodium periodate (NaIO4). The
mixture was stirred vigorously for 15 min. To this biphasic solution, 12.2 mg (0.06 mmol) of RuCl3 · 3H2O was
added, and the mixture was stirred vigorously for 6 h at room
temperature. Then, 24.4 ml of dichloromethane was added, and the phases
were separated. The upper aqueous layer was extracted three times with
dichloromethane. The combined organic extracts were dried over
anhydrous MgSO4 and concentrated. The residue was purified
by column chromatography (the eluent was ethyl acetate with a few drops
of acetic acid until the elution of the aldehyde and then methanol).
The yield was 28%, with 1H NMR (400.13 MHz,
CDCl3) ppm 1.45 (s, 9H), 3.35 (t, 2H, J = 5 Hz), 3.65 (t, 2H, J = 5 Hz), 4.10 (s, 2H), 5.30 (s,
1H), 8.20 (s, 1H).
(iii) Deprotection of the amino group.
The amino group was
deprotected as described previously (26). A suspension of 40 mg (0.18 mmol) of the N-protected acid in 10 ml of a 3 N HCl solution
in ethyl acetate was stirred vigorously at room temperature for 30 min.
The solution was removed in vacuo, and the oil was triturated with
ether (3 ml). The ether layer was removed, and the aqueous phase was
evaporated. After the sample was dried overnight, the pure chlorhydrate
of 2-(2-aminoethoxy)acetate was obtained with a yield of 85%, with
1H NMR (400.13 MHz, D2O) ppm 3.97 (s, 1H),
3.69 (t, 2H, J = 3.6 Hz), 3.07 (t, 2H, J = 3.6 Hz).
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RESULTS |
Degradation of morpholine by M. aurum MO1.
Resting
M. aurum MO1 cells (5 g of wet cells in 50 ml of Knapp
buffer) were incubated with 10 mM morpholine at 30°C with agitation (200 rpm) for 72 h.
In order to find the best conditions for complete morpholine
degradation, the first experiments were carried out with different [morpholine]/[bacterium] ratios. The amount of bacteria was varied from 100 to 10 g of wet cells in 1 liter of Knapp buffer, while the morpholine concentration was kept constant (10 mM [0.87 g · liter
1]). In all cases, the same intermediates were
detected in 1H-NMR spectra. However, when the cell
concentration was low (10 g · liter1), the
degradation of morpholine stopped before completion. This could result
from the inhibitory effect of ammonia resulting from the degradation of
morpholine, as Mazure and Truffaut (19) showed that ammonia
was toxic to this strain. The best results were obtained with a cell
concentration of 100 g · liter1. The following
experiments were carried out with this concentration.
Samples (1 ml) were taken every hour, and the supernatants (500 µl)
were analyzed by
1H-NMR spectroscopy after adjustment of
the pH to 10 to avoid changes
in chemical shifts. The spectra obtained
were compared with those
of the negative controls (incubation of cells
under the same conditions
in the absence of substrate and incubation of
the substrate in
the buffer without cells). Spectra collected at 0, 10, and 20
h are presented in Fig.
2A.
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FIG. 2.
(A) Kinetics of morpholine degradation by M. aurum MO1. Resting cells (5 g of wet cells in 50 ml of Knapp
buffer [KH2PO4, 1 g · liter1; K2HPO4, 1 g · liter1; FeCl3, 4 mg · liter1; MgSO4 · 7H2O, 40 mg · liter1; pH 6.6]) were incubated with 10 mM
morpholine at 30°C with agitation (200 rpm) for 72 h. Samples (1 ml) were collected every hour for 12 h and from time to time until
72 h; after centrifugation, the supernatants of these samples were
analyzed by 1H-NMR spectroscopy at 300.13 MHz.
TSPd4 was used as a reference for chemical shifts and
quantification. (B). Expanded scale, from 2.60 to 3.98 ppm, of the 10-h
spectrum. M, morpholine; G, glycolate; Y, 2-(2-aminoethoxy)acetate.
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In the spectrum obtained at time zero, three main signals are visible:
a singlet at 0 ppm that belongs to the methyl groups
of
TSPd
4 and two pseudotriplets at 2.88 and 3.72 ppm that
correspond
to CH
2(b) and CH
2(a) of morpholine
(Fig.
1). The singlet corresponding
to the NH of morpholine was not
detected in water because of the
quadrupolar moment of
14N.
At 10 h (Fig.
2B), the signals of morpholine were decreasing
while
a singlet at 3.95 ppm was increasing. This signal was assigned
to
glycolate as evidenced by addition of the commercial compound
to the
sample. Three new signals were also present, resonating
at 3.96 ppm
(singlet), 3.67 ppm (pseudotriplet), and 3.05 ppm
(pseudotriplet).
These signals are compatible with those of
2-(2-aminoethoxy)acetate
CH
2(c), CH
2(a), and
CH
2(b), respectively (Fig.
1). At 20 h,
morpholine
was exhausted and all the intermediate compounds were
decreasing
and had almost disappeared.
In order to confirm the assignment for the intermediate,
2-(2-aminoethoxy)acetate was synthesized. Vieles and Séguin
(
32)
first described the synthesis of this compound, but the
yield
obtained was too low to be used. Therefore, we developed a new
synthesis strategy in three steps (Fig.
3). The first step corresponded
to the
protection of the amino function with
di-
tert-butyldicarbonate
in order to avoid the reaction of
this function during the oxidation.
This reaction was almost
quantitative (yield, 92%). For the next
step, which was oxidation of
the alcohol into acid, the choice
of the oxidant was more difficult,
since it should not be too
acidic or too strong in order not to remove
the N protection.
RuCl
3 · 3H
2O was
chosen (
2). A mixture of the aldehyde and
the acid was
obtained. The most difficult point was the purification
of this acid;
column chromatography on silica gel with a polar
eluent was performed.
The last step was the deprotection of the
amino group. The N-protected
acid was placed in a 3 N HCl solution
in ethyl acetate, and the
chlorhydrate of 2-(2-aminoethoxy)acetic
acid was obtained with a good
yield of 85%. After adjustment of
the pH to 10, 2-(2-aminoethoxy)acetate was added to the sample
collected at 12 h. The resonances were perfectly overlapping (data
not shown).
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FIG. 3.
Synthesis of the chlorhydrate of
2-(2-aminoethoxy)acetate. Boc2O,
di-tert-butyldicarbonate; EtOAc, ethyl acetate.
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Quantitative analysis of the kinetics of degradation of morpholine was
performed by integrating the signals of the different
metabolites in
1H-NMR spectra; the measured areas were compared to the
integral
of the TSPd
4 signal. The different concentrations
of metabolites
were calculated from these integrals as described in
Materials
and Methods. Under these conditions, the limit of
quantification
for an individual metabolite was estimated to be 0.05 mM. Figure
4 shows one example of the
time courses for the concentrations
of morpholine, glycolate, and
2-(2-aminoethoxy)acetate. The kinetics
were quite reproducible,
as shown in the inset, where data from
five independent experiments are
gathered.
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FIG. 4.
Time courses for the concentrations of morpholine (×),
glycolate ( ), and 2-(2-aminoethoxy)acetate ( ) during the
degradation of morpholine (10 mM) by M. aurum MO1 cells at
100 g · liter1. The quantification was made by
integrating the signals in 1H-NMR spectra relative to the
area for the reference TSPd4 (see Fig. 2). (Inset) Data
from five independant experiments concerning the degradation of
morpholine; a polynomial calculation was used to plot the final
curve.
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Morpholine was almost exhausted after 10 h of incubation. Its rate
of degradation was about 0.85 mM/h. Glycolate and
2-(2-aminoethoxy)acetate
concentrations increased with time until 10 to
12 h and then decreased.
No more glycolate was detected after
20 h or even longer. Consequently,
glycolate is likely to be
degraded in the cells.
Degradation of ethanolamine and glycolic acid by M. aurum MO1.
Mazure and Truffaut (19) showed that
M. aurum MO1 was able to grow on ethanolamine in the absence
of morpholine. In contrast, under the same conditions, no growth was
observed on glycolic acid. This prompted us to monitor the kinetics of
degradation of these two compounds, which are thought to be
intermediates in the morpholine biodegradation pathway (Fig. 1). The
most efficient condition for morpholine degradation, i.e., 100 g
of bacteria · liter1, was used, with 10 mM
ethanolamine or 13 mM glycolate.
An example of a
1H-NMR spectrum collected after 5 h of
incubation with ethanolamine is shown in Fig.
5A. The triplets resonating
at 3.02 and
3.75 ppm correspond, respectively, to CH
2(b) and
CH
2(a)
of ethanolamine (Fig.
1). A resonance at 2.01 ppm
(singlet) was
assigned to acetate after addition of the commercial
compound.
The resonance at 3.83 ppm was also present in the control
sample
and thus is not a metabolite from morpholine degradation. The
time courses for the metabolite concentrations are presented in
Fig.
5B. Ethanolamine was degraded in 8 h at a rate of about
1.25
mM/h. During this degradation, the acetate level remained
very
low (about 0.3 mM).
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FIG. 5.
Kinetics of degradation of ethanolamine (10 mM) by
M. aurum MO1 (100 g · liter1). The
conditions were as described for Fig. 2. (A) 1H-NMR
spectrum collected after 5 h of incubation. (B) Time courses for
the concentrations of ethanolamine (×) and acetate ().
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As shown before, glycolate has been identified as one of the
intermediates in the morpholine degradation pathway. Study of
its
degradation kinetics is therefore particularly interesting
(Fig.
6). When glycolic acid was added directly
to the flask,
under the usual conditions, the degradation was
rapidly stopped
(in about 4 h), whereas the
1H-NMR
spectrum analysis of the morpholine degradation showed an
increase and
then a total disappearance of glycolic acid in 20
h. The pH was
checked and was found to be very acidic (pH 3).
Cells can no longer
degrade glycolic acid at such a pH. The pH
of the culture medium was
then adjusted to 7 with various bases
(KOH, NH
4OH, and
morpholine) before the addition of the bacteria.
Under these
conditions, the degradation of glycolate was complete
in 4 h, no
matter which base was used. The rate of degradation
was very high, 3.25 mM/h.
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FIG. 6.
Kinetics of degradation of glycolic acid (13 mM) by
M. aurum MO1 (100 g · liter1) without
pH adjustment ( ) and with adjustment of the pH to 7 by KOH (×),
NH4OH (), and morpholine ().
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DISCUSSION |
In this paper, 1H-NMR spectroscopy, performed directly
on the incubation medium, was shown to be a useful tool to study the degradation of morpholine by M. aurum MO1. This approach was
useful to directly quantify the degradation of morpholine. The NMR
spectra collected at different incubation times showed that this strain is able to degrade morpholine at a rate of 0.85 mM/h under our conditions. It is worth noting that biodegradation began immediately; no latency period was observed. This observation shows that either no
induction existed or the induction time was very short (less than
1 h).
The 1H-NMR technique also allowed us to identify
unambiguously, for the first time, glycolate and
2-(2-aminoethoxy)acetate as intermediates of the biodegradation
pathway. This suggests that M. aurum MO1 cleaved the C-N
bond of the morpholine ring. These intermediates were suggested by
Swain et al. (28) in the case of M. chelonae MorG
but were never directly evidenced. In our experiments we did not find
evidence of intermediates of the ethanolamine branch
(acetaldehyde or acetate) during incubation with morpholine. However,
we have shown that M. aurum cells could degrade ethanolamine
through the classical pathway via acetate. This confirms the
results of Mazure and Truffaut (19) concerning growth on
this substrate without induction of morpholine. It is interesting that
the rate of degradation of ethanolamine measured under our conditions
(1.25 mM/h) was threefold higher than that of morpholine (0.85 mM/h).
Glycolate has been identified as the major intermediate of the
morpholine degradation pathway. Although M. aurum MO1 was
not able to grow on glycolic acid (19), we have shown that
glycolic acid was degraded very rapidly by this strain (about 3.25 mM/hour). This degradation is pH dependent and can occur only if the pH is not too acidic. The nature of the base used for pH adjustment is not
important. Mazure and Truffaut (19) reported that no growth
was detected in the absence of morpholine with glycolic acid at pH 6.5 but that the presence of 0.1 g of morpholine · liter1 induced degradation of this acid. Our results are
different, showing a complete degradation of glycolate in the absence
of morpholine (assays with KOH or NH4OH). Therefore, the
enzymes required for this degradation are present in the microorganism, and no induction by morpholine is necessary.
In conclusion, this work is a pioneer in the direct quantification and
identification of the morpholine degradation pathway of M. aurum MO1. The use of in situ 1H-NMR spectroscopy
allows direct determination of the intermediates formed. Work is now in
progress to identify other metabolites (particularly by analyzing the
intracellular medium) and to understand the regulation of this pathway.
The methodology described here, using 1H-NMR spectroscopy,
is general and easy to apply and could be used to investigate many biodegradative processes. This technique was used in the following companion paper (24) to study the morpholine biodegradation pathway with another strain. In this second article, we show evidence for the involvement of a cytochrome P-450 in morpholine degradation.
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ACKNOWLEDGMENTS |
This work was supported by interdisciplinary programs of CNRS
"PIRSEM" and "ECOTECH."
We acknowledge Anne-Lise Etienne, coordinator of the CNRS programs. We
greatfully acknowledge Christelle Roland and Tran Thi Minh Ngoc for
their help. We thank J. S. Cech for the gift of M. aurum MO1.
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FOOTNOTES |
*
Corresponding author. Mailing address: Laboratoire de
Synthèse, Electrosynthèse et Etude de Systèmes
à Intérêt Biologique, UMR 6504 CNRS, Université
Blaise Pascal, 63177 Aubière Cedex, France. Phone: 33 4 73 40 77 14. Fax: 33 4 73 40 77 17. E-mail: amdelort{at}chimtp.univ-bpclermont.fr.
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Appl Environ Microbiol, January 1998, p. 153-158, Vol. 64, No. 1
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Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
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