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Applied and Environmental Microbiology, May 2000, p. 1877-1882, Vol. 66, No. 5
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Toluene Monooxygenase-Catalyzed Epoxidation
of Alkenes
Kevin
McClay,1
Brian G.
Fox,2 and
Robert J.
Steffan1,*
Envirogen, Inc., Lawrenceville, New Jersey
08648,1 and Department of Biochemistry,
College of Agricultural and Life Sciences, University of Wisconsin,
Madison, Wisconsin 537052
Received 30 August 1999/Accepted 20 February 2000
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ABSTRACT |
Several toluene monooxygenase-producing organisms were tested for
their ability to oxidize linear alkenes and chloroalkenes three to
eight carbons long. Each of the wild-type organisms degraded all of the
alkenes that were tested. Epoxides were produced during the oxidation
of butene, butadiene, and pentene but not hexene or octadiene. A strain
of Escherichia coli expressing the cloned toluene-4-monooxygenase (T4MO) of Pseudomonas mendocina KR1
was able to oxidize butene, butadiene, pentene, and hexene but not octadiene, producing epoxides from all of the substrates that were
oxidized. A T4MO-deficient variant of P. mendocina KR1
oxidized alkenes that were five to eight carbons long, but no epoxides were detected, suggesting the presence of multiple alkene-degrading enzymes in this organism. The alkene oxidation rates varied widely (ranging from 0.01 to 0.33 µmol of substrate/min/mg of cell protein) and were specific for each organism-substrate pair. The enantiomeric purity of the epoxide products also varied widely, ranging from 54 to
>90% of a single epoxide enantiomer. In the absence of more preferred
substrates, such as toluene or alkenes, the epoxides underwent further
toluene monooxygenase-catalyzed transformations, forming products that
were not identified.
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INTRODUCTION |
The reactivity of epoxides makes
them useful and important intermediates for industrial chemical
syntheses, including the production of pharmaceuticals, agrochemicals,
and polymers. Although there are many applications for racemic
epoxides, which are usually produced by chemical means, the demand for
the production of enantiomerically pure feedstocks, including epoxides,
has increased in recent years. Between 1996 and 1997, the sale of
enantiomerically pure pharmaceuticals increased 21%, and similar
increases are expected in the agrochemical and polymer markets
(19). This increased interest in enantiomeric purity arises
from the observation that the (R) and (S)
enantiomers of an individual compound often have appreciably different
biological and physical properties. For example, the (S)
enantiomer of carvone gives caraway seeds their distinctive odor,
whereas the (R) enantiomer is perceived as spearmint
(10). Similarly, (S)-thalidomide can cause severe
birth defects, while (R)-thalidomide is a safe and effective
sedative (10). These dramatic differences have led the U.S.
Food and Drug Administration to require that each enantiomer of a
racemic drug be tested individually prior to approval of a drug
(10), thereby greatly increasing the cost of bringing a new
drug to market. By using enantiomerically pure intermediates in drug
synthesis, the number of isomers of a new drug that require testing
prior to approval can be reduced.
One possible method for producing enantiomerically pure epoxides, that
of using enzymes to enantioselectively insert oxygen across the carbon
double bonds of alkenes, has been explored, with various degrees of
success. The styrene monooxygenase of Pseudomonas sp. strain
VLB120 converts styrene to (S)-styrene oxide with an
enantiomer excess of >99% (16). Another enzyme, the alkane
hydroxylase of Pseudomonas oleovorans, converts
1,7-octadiene to optically active (R)-7,8-epoxy-1-octene
with an enantiomeric purity of 92%; the latter is also oxidized by the
enzyme, forming (R,R)-1,2-7,8-diepoxyoctane with
an enantiomeric purity of 83% (12). These observations led
to industrial applications for the synthesis of the drugs Metoprolol
and Atenolol (1).
A class of oxygenases with a non-heme diiron cluster at the catalytic
center is also known to form epoxides from alkenes. One such
enzyme, the soluble methane monooxygenase (MMO) of
Methylosinus trichosporium OB3b, mediated the epoxidation of
propene, 1-butene, 2-butene, and 1,3-butadiene, but with very low
enantiomeric specificity (less than 64% of a single isomer)
(15). The epoxides formed from propene, butene, and
butadiene were not oxidized further by MMO under the experimental
conditions described, but in other experiments, ethylene epoxide and
cis-dichloroethylene (cis-DCE) epoxide were shown
to be substrates for MMO (20). MMO was unable to oxidize the
larger alkenes 1-pentene, cyclohexene, and 3-methyl-butene (15). In contrast, the alkene monooxygenase of
Xanthobacter sp. strain Py2 can form epoxides from alkenes
and chlorinated alkenes, but with a higher degree of enantiomeric
selectivity. For example, the oxidation of 3-chloropropene yielded 80%
(S)-3-chloro-1,2-epoxypropene, whereas the oxidation of
1-butene yielded 94% of the (R) isomer (6, 7).
The formation of epoxides by other bacterial systems has been reviewed
in detail elsewhere (1, 2).
In this report, we discuss the epoxidation of alkenes by a subgroup of
the non-heme diiron-containing enzymes, the toluene monooxygenases,
which hydroxylate the aromatic ring of toluene, forming
ortho-, meta-, and para-cresols. The
rate and the enantiomeric selectivity of alkene oxidations varied with
each enzyme-substrate pair. In some cases, the enantiomeric selectivity
of the epoxidation reactions was >90%. We also found evidence that
the epoxides formed as a result of alkene oxidation underwent
additional toluene monooxygenase-mediated transformations.
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MATERIALS AND METHODS |
Chemicals.
The chemicals 1-butene (99%), 2-butene (99%,
cis-trans mixture), 1,3-butadiene (99%), epoxybutane
(99%), 1,3-butadiene monoepoxide (BME) (95%), butadiene diepoxide
(97%), epichlorohydrin (99%), 1-pentene (99%), 2-pentene (99%,
cis-trans mixture), hexene (97%), octadiene (95%),
2-chloropropene (99%), 2,3-dichloropropene (95%), 1,2-butanediol
(95%), toluene (95%), triethylamine,
4-(p-nitrobenzyl)pyridine (PNBP), ethylene glycol, and
isopropyl-
-thiogalactopyranoside (IPTG) were obtained from Aldrich
Chemicals (Milwaukee, Wis.).
Growth and preparation of cells.
Pseudomonas
mendocina KR1 (22), P. mendocina ENVpmx1 (a
toluene-4-monooxygenase [T4MO]-deficient mutant of KR1) (K. McClay and R. J. Steffan, Abstr. 97th Gen. Meet. Am. Soc. Microbiol., abstr. K36, p. 348, 1997), Ralstonia pickettii PKO1
(3), Burkholderia cepacia G4 (18),
Burkholderia sp. strain ENVBF1 (13), and Pseudomonas sp. strain ENVPC5 (13) were cultured
overnight at 30°C in shake flasks containing basal salts medium
(8) supplemented with 0.4% sodium glutamate. Toluene was
included in the vapor phase of the cultures when the induction of
toluene oxygenases was desired. Prior to substrate degradation assays,
the cultures were harvested by centrifugation and resuspended in basal
salts medium to an optical density at 550 nm (OD550) of 2, unless otherwise indicated. A standard curve of optical density versus
protein concentration for each strain was used to calculate the amount of protein per milliliter of the resuspended cultures.
Escherichia coli DH10B containing plasmid pRS202
(17) was prepared in a similar manner, except that the
strain was grown at 37°C in Luria-Bertani medium and was resuspended
to an OD550 in Luria-Bertani medium of 4 with 0.3 to 1 mM
IPTG to induce the expression of T4MO.
Substrate degradation assays.
To determine the substrate
range and the rates of alkene oxidation for the various toluene
monooxygenase-producing organisms, duplicate 5-ml aliquots of the
resuspended cultures were dispensed into 25-ml serum vials and crimp
sealed with Teflon-faced septa. Gaseous substrates were added as pure
compounds using a gas-tight syringe, and other substrates were added
from 20% stock solutions in dimethylformamide (Fig.
1). The amounts of individual substrates added were as follows, unless otherwise indicated: 1-butene, 2-butene, and 1,3-butadiene, 2.2 µM; epoxybutane, BME, and epichlorohydrin, 8 µM; 1- and 2-pentene, 9 µM; hexene, 4 µM; octadiene, 4.5 µM; 2-chloropropene, 11 µM; and 2,3-dichloropropene, 12 µM. The serum vials were then placed horizontally on a rotary shaker operating at 100 rpm. During incubation, the temperature was maintained at between 20 and 22°C for the pseudomonads and at 37°C for E. coli.
The serum vials were periodically removed from the shaker, and a 10- to
25-µl portion of the headspace gas was withdrawn through the septa
and injected into a gas chromatograph (GC) equipped with a 30-m Vocol
column (Supelco Inc., Bellefonte, Pa.) maintained at 160°C and a
flame ionization detector. This procedure allowed us to monitor the
concentrations of both the alkene substrates and the epoxide products.
The same protocol was used to detect the formation of 1,2-butanediol
and butadiene diepoxide, except that 1 µl of culture medium was
injected into the column instead of headspace gas.
Verification of epoxide formation.
The commercially
available epoxides butene epoxide, BME, and butadiene diepoxide were
used in GC analyses to determine the retention times of the authentic
compounds and to quantitate the conversion of the alkenes to the
corresponding epoxides. A modification of the method of van Hylckama et
al. (20) was used to verify that the observed peaks that
coeluted with the authentic compounds were volatile epoxides; the
epoxides were conjugated with PNBP to form intensely colored adducts of
the epoxides. Serum vials were prepared as described above, except that
prior to sealing, a glass test tube (5 by 50 mm) was placed inside each
vial with the opening of the test tube extending out of the liquid. The substrate was then added, and the vials were incubated with moderate shaking at a 45° angle, preventing the culture liquid from entering the test tube. The transformation of the alkenes was monitored by GC
analyses. After the rate of transformation decreased significantly or
90% of the alkene substrate was depleted from the headspace, 400 µl
of 100 mM PNBP dissolved in ethylene glycol was injected through the
septum into the open end of the test tube. The vials were then
incubated for 5 h before they were opened, and the epoxide-binding PNBP was withdrawn. The total volume of PNBP solution was combined with
an equal volume of acetone-triethylamine (50:50) and mixed rapidly.
Epoxide-PNBP adducts were detected by spectral analysis at 400 to 700 nm.
Determining enantiomeric ratios.
To determine the ratio of
(R) and (S) isomers of the epoxides formed from
the oxidation of the alkenes, samples were analyzed for the presence of
the epoxides and the alkenes by GC analyses as described above. When
the epoxide concentration neared its maximum or when 80 to 90% of the
substrate was oxidized, the enantiomeric ratio of epoxides produced was
determined by injecting a sample of the headspace gas into a GC
equipped with a chiral separation column (RT-BDEXSE; Restek, Inc.,
Bellefonte, Pa.) and a flame ionization detector. The column was
maintained at 50°C.
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RESULTS |
Alkene oxidation and identification of oxidation products.
All
of the wild-type toluene monooxygenase-producing organisms tested were
able to oxidize alkenes. Greater than 95% of the added butadiene (2.2 µmol) was oxidized by strains G4 and ENVBF1 during the first 5 h
of incubation, whereas strains KR1 and ENVPC5 oxidized only 50% of the
butadiene in 20 h (Fig. 2). Greater
than 95% of the added 2-butene (2.2 µmol) was oxidized by all the
strains tested, except ENVpmx1, within 20 h, with strains KR1 and
ENVPC5 having higher initial degradation rates than strains G4 and
ENVBF1 (Table 1). With the T4MO-deficient
strain ENVpmx1, the concentrations of butadiene and 2-butene decreased
by less than 10% during 20-h incubations. The T4MO clone, E. coli pRS202, oxidized all of the butenes tested.
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FIG. 2.
Oxidation of 1,3-butadiene by toluene
monooxygenase-producing organisms. Each datum point represents the
average of the analysis of duplicate sample vials, with the range
indicted by error bars. Symbols: closed circle, P. mendocina
KR1; open triangle, ENVpmx1; open square, pRS202; closed triangle,
ENVPC5; closed square, ENVBF1; closed diamond, B. cepacia
G4.
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1-Pentene and 2-pentene were oxidized by all of the wild-type strains
tested. In the case of the T4MO knockout mutant ENVpmx1,
an extended
lag period was followed by the rapid transformation
of 2-pentene that
exceeded the rates of 2-pentene oxidation observed
in the wild-type
organisms (Fig.
3). Similarly, hexene and
octadiene
were also transformed more rapidly by strain ENVpmx1 than by
the
wild-type organisms.
E. coli(pRS202) expressing T4MO
oxidized
1-pentene, 2-pentene, and hexene but did not oxidize
octadiene.
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FIG. 3.
Oxidation of 2-pentene by toluene
monooxygenase-producing organisms. Each datum point represents the
average of the analysis of duplicate sample vials, with the range
indicted by error bars. Symbols: closed circle, P. mendocina
KR1; open triangle, ENVpmx1; open square, pRS202; closed triangle,
ENVPC5; closed square, ENVBF1; closed diamond, B. cepacia
G4.
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The chlorinated alkene 2-chloropropene was oxidized by strains KR1,
ENVPC5, and ENVBF1. Even though all three of these organisms
appear to
produce the same class of toluene monooxygenase (as
indicated by the
appearance of
para-cresol in the culture medium
when grown
on toluene), there were differences in their ability
to oxidize this
compound. Strains KR1 and ENVPC5 oxidized 2.1
and 4 µmol of substrate
in the first 3.5 h of incubation, respectively,
and oxidized only
an additional 0.54 and 1.0 µmol, respectively,
during the following
17 h. Strain ENVBF1 oxidized 2.3 µmol of
2-chloropropene in the
first 3 h of incubation and oxidized 6.3
µmol of substrate in
the following 17 h. It is not known why ENVBF1
continued to
oxidize 2-chloropropene after oxidation by the other
two T4MO-producing
organisms had
ceased.
The ability of the T4MO-producing organisms to oxidize alkenes was
adversely affected by the presence of an additional chlorine
atom on
the substrate molecules. The amounts of 2,3-chloropropene
oxidized by
strains KR1, ENVPC5, and ENVBF1 were 95.2, 66.0, and
75.6% smaller
than the amounts of 2-chloropropene oxidized by
the same strains,
respectively. Strain G4 was not tested on 2-chloropropene,
but it was
able to oxidize twice as much 2,3-chloropropene (6.7
µmol) as any of
the T4MO-expressing organisms in this
assay.
The amounts of the 3- and 4-carbon alkenes oxidized by the cultures
were related to the specific activity of the organisms
toward the given
alkene. For example, 1,3-butadiene was oxidized
by strain G4 at an
initial rate of 0.19 µmol/min/mg of cell protein
(Table
1) and was
oxidized to a concentration below the limits
of detection within the
first 6 h of incubation (Fig.
2). Strain
KR1 had an initial
oxidation rate of 0.07 µmol/min/mg of cell
protein and ultimately
oxidized only 1.1 µmol (50%) of butadiene
in 20 h. With
2-butene as a substrate, KR1 had an initial oxidation
rate of 0.26 µmol/min/mg of cell protein and oxidized all of the
2-butene (2.2 µmol) in 3.5 h. Strain G4 oxidized 2-butene at an
initial
oxidation rate of only 0.14 µmol/min/mg of cell protein
and required
20 h to oxidize >95% of the added
compound.
The extent of pentene and halogenated propene oxidation achieved by the
individual organisms was not well correlated with
the initial rates of
oxidation of the compounds. Strain KR1 had
the highest initial
2-pentene oxidation rate (0.33 µmol/min/mg
of cell protein). This
rate was more than twice the initial rate
of 2-pentene oxidation by G4
and BF1 (Table
1), yet BF1 and G4
oxidized more 2-pentene than KR1
during a 20-h incubation (Fig.
3). The T4MO-deficient mutant of KR1,
strain ENVpmx1, oxidized
5- to 8-carbon alkenes efficiently, apparently
utilizing an alternate
enzyme system. Strain ENVPC5 had a higher
initial oxidation rate
than ENVBF1 for 2- and 2,3-chloropropene (Table
1) but ultimately
oxidized less of these substrates over a 20-h
incubation (data
not
shown).
Epoxide formation and degradation.
GC analysis showed that
during 1,3-butadiene, 2-butene, 1-pentene, and 2-pentene oxidation by
the wild-type organisms and E. coli(pRS202), a transient
secondary peak appeared on chromatograms. A similar peak was observed
during the oxidation of hexene by pRS202 but not when the wild-type
organisms oxidized this substrate. These secondary peaks increased in
proportion to the amount of alkene oxidized and then decreased
following the depletion of the alkene. The peak formed during the
oxidation of 1-butene and butadiene coeluted with the commercially
available butene epoxide and BME. To verify that this peak and the
corresponding peaks formed during the oxidation of the other alkenes
were epoxides, they were conjugated with PNBP as described in Materials
and Methods. Spectral analysis showed that the PNBP conjugates of the
products of 1-butene and butadiene oxidation had absorbance maxima
identical to those of the conjugates of the purchased epoxides and
agreed closely with the data obtained for other epoxides (4,
20). The conjugates of the pentenes and 2-butene had similar
absorbance spectra. From these data it was concluded that the secondary
peaks were epoxides.
Even though the T4MO-deficient mutant ENVpmx1 efficiently oxidized
pentene, hexene, and octadiene, no epoxides were detected
during the
oxidation of any of these compounds by this strain.
Similarly, no
epoxides were detected during the oxidation of hexene,
octadiene,
2-chloropropene, or 2,3-chloropropene by the wild-type
organisms.
The stoichiometry of epoxide formation was evaluated by incubating
toluene-induced G4 with 4.4 µmol of 1,3-butadiene and monitoring
both
butadiene and BME concentrations. Initially, there was a
nearly
stoichiometric conversion of butadiene to BME (>95%), followed
by a
decrease in BME concentration after the parental compound
was depleted
(Fig.
4). In contrast, when G4 was
incubated with
2.2 µmol of 2-butene, only 43% of the 2-butene
oxidized could
be detected as the epoxide product. Efforts to detect
2-butene-1-ol
in liquid media were not successful.
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FIG. 4.
Stoichiometric conversion of 1,3-butadiene to
monoepoxide by T2MO-expressing B. cepacia G4. Each datum
point represents the average of the analysis of duplicate sample vials,
with the range indicated by error bars. Symbols: closed circle,
1,3-butadiene; open circle, BME.
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To determine if the disappearance of the epoxides was caused by
chemical or enzymatic reactions, BME was incubated with both
toluene-induced and uninduced cultures of strain G4. Induced cells
incubated with pure BME oxidized 8 µmol of the substrate in the
first
50 min of incubation, whereas uninduced cells of G4 oxidized
less than
1 µmol in the same time period. When toluene-induced
cells of G4 were
incubated in the presence of both toluene and
BME, they degraded 0.5 µmol of BME in 50 min (Fig.
5). A
similar
inhibition of BME transformation was observed when BME was
coincubated
with butadiene. The oxidation of both toluene and butadiene
was
unaffected by the presence of BME, but butadiene oxidation was
inhibited by toluene (data not shown). Similar results were obtained
with ENVBF1. The uninduced cells degraded less than 1.3 µmol of
BME
in 24 h, whereas the induced cells oxidized 13.4 µmol.
Epichlorohydrin
and butene epoxide also served as substrates for
toluene monooxygenases
(Table
2).
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FIG. 5.
Inhibition of butadiene and BME transformation by
toluene in B. cepacia G4. Each datum point represents the
average of the analysis of duplicate sample vials, with the range
indicated with error bars. Symbols: closed circle, butadiene; open
circle, butadiene in the presence of toluene; closed square, BME; open
square, monoepoxide in the presence of toluene. The initial
concentration of toluene in samples was 20 µmol. When the quantity of
toluene was reduced to 10 to 11 µmol, an additional 10 µmol of
toluene was added through the septa.
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Enantiomeric ratios of biologically produced epoxides.
The
epoxides formed from the oxidation of 1-pentene and 1,3-butadiene were
analyzed by chiral chromatography, and the results of these analyses
are presented in Table 3. The
enantiomeric selectivity observed during the formation of BME from
1,3-butadiene by the various toluene monooxygenases differed, but all
of the oxygenases tested favored the production of the
(S)-enantiomer. The highest selectivity occurred with strain
G4 (toluene-2-monooxygenase [T2MO]) [91.9% (S)], and
the lowest selectivity occurred with strain ENVPC5 (T4MO) [67.3%
(S)].
The oxidation of 1-pentene showed greater variation in the observed
selectivity, but in each case, a larger percentage of
the epoxide
formed was of the (
R)-enantiomer. The product distribution
ranged from 100% (
R)-enantiomer formed by strain G4 to a
low of
54.2% (
R)-enantiomer formed by strain KR1 (Table
3).
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DISCUSSION |
Several non-heme diiron-containing monooxygenases form epoxides as
the primary product of halogenated ethene oxidation (4, 21).
In this study, we report the oxidation of 4-, 5-, and 6-carbon alkenes
to their corresponding epoxides by several toluene monooxygenases. The
enzymes tested oxidize toluene at the ortho,
meta, or para position; each has considerable
amino acid sequence homology to the hydroxylases of MMO, the alkene
monooxygenase of Xanthobacter sp. strain PY2, and the alkene
monooxygenase of Rhodococcus rhodochrous B276
(23), all of which are known to oxidize alkenes to their corresponding epoxides. We also screened the toluene dioxygenases of
Pseudomonas putida F1 (24) and
Pseudomonas sp. strain GZ-9 and found that although they
oxidized alkenes, no epoxides were formed (data not shown). These
findings are consistent with those of Lange and Wackett
(11), who showed that the toluene dioxygenase of P. putida F1 oxidizes a wide range of alkenes and haloalkenes, forming alcohols, diols, and ketones.
All of the toluene monooxygenase-producing strains that we examined
catalyzed the epoxidation of short-chain alkenes. When expressed in
E. coli, the T4MO of KR1 formed epoxides from 1-butene, 2-butene, 1,3-butadiene, 1-pentene, 2-pentene, and 1-hexene, but octadiene was not oxidized. The wild-type organisms R. pickettii PKO1, B. cepacia G4, P. mendocina
KR1, Pseudomonas sp. strain ENVBF1, and
Pseudomonas sp. strain ENVPC5 oxidized all of the nonhalogenated alkenes tested. However, the oxidation of hexene and
octadiene by the wild-type organisms did not lead to the formation of epoxides.
Although 2-chloropropene and 2,3-chloropropene were oxidized by the
strains tested, no epoxides were detected by GC analysis. Nonetheless,
we suspect that epoxides were formed. The epoxides of 2-chloropropene
and 2,3-chloropropene would have a chlorine atom bonded to one of the
epoxide ring carbons, and such an arrangement would result in an
unstable epoxide that would undergo rapid chemical hydrolysis, thereby
preventing accumulation and detection. Similar results have been
described for the oxidation of various halogenated alkenes, such as
trichloroethylene (TCE) and vinyl chloride (4, 20).
The rates of alkene oxidation observed during this study varied between
the substrates and the strains tested, yet there was no apparent
correlation between the regiospecificity of toluene oxidation and the
rate of alkene oxidation by these enzymes. However, because whole-cell
assays were used in this study, the oxidation rates observed could have
been affected by factors other than the kinetic characteristics of the
toluene monooxygenases. For example, differences in membrane
compositions or transport mechanisms or the presence of alternative
alkene oxidation pathways could affect the observed transformation
rates. This situation was demonstrated by comparing the rates of alkene
degradation by P. mendocina KR1, the T4MO-deficient mutant
ENVpmx1, and E. coli(pRS202) expressing T4MO. Essentially no
1,3-butadiene or 2-butene was oxidized by ENVpmx1 in the first 5 h
of incubation, whereas KR1 and E. coli(pRS202) oxidized 30 to 77% of the butadiene and nearly 100% of the 2-butene. Because T4MO
was shown to facilitate short-chain alkene degradation in KR1, these
results were not surprising. However, ENVpmx1 oxidized 2-pentene,
hexene, and octadiene more rapidly than either E. coli(pRS202) or KR1. Although we previously have shown that strain
KR1 can oxidize alkanes (13), presumably by using a separate
alkane oxidation pathway, it was surprising that the T4MO-deficient
mutant would oxidize a greater quantity of the alkenes than the
wild-type strain, which has both functional pathways available to
oxidize alkenes. However, ENVpmx1 does contain the lux
operon, which KR1 lacks. The light-producing apparatus of the
lux operon catalyzes the oxidation of aldehydes
(5), and it is possible that an enzyme present in KR1
converts pentene, hexene, and octadiene to aldehydes, which are then
rapidly utilized by the lux genes, thereby participating in
the metabolism of the alkenes. We observed that the alkenes used in
this study induced the expression of the T4MO::lux
transcriptional fusion present in ENVpmx1 (data not shown), ensuring
that the products of the lux genes would be present
throughout the assay.
Another possible explanation for the difference in observed alkene
oxidation rates between these two strains is that the epoxides that
result from alkene oxidations damage T4MO-producing cells. It has been
reported that TCE and vinyl chloride epoxides are harmful to cells,
presumably because they react with cellular components (4,
14). Even though unhalogenated epoxides are more stable than
halogenated ethenes, with a half-life on the order of days rather than
seconds (20), it is still possible that they have a similar
toxic effect. If ENVpmx1 does not produce epoxides during alkene
oxidation, the overall health of the cells may be better than that of
toluene monooxygenase-expressing cultures, allowing faster degradation
of the alkenes.
Yeager et al. (21) recently reported that the T2MO of G4
produces epoxides during the oxidation of ethene and propene, with 96%
of the propene being converted to propene oxide. We obtained similar
results, with 104% (±10%) of the butadiene being converted to BME.
When 2-butene was oxidized by G4, however, the epoxide did not
accumulate stoichiometrically. Although all of the added 2-butene was
depleted, only 43% was recovered as the corresponding epoxide, and the
remaining 57% was unaccounted for (data not shown). Similarly, when
MMO oxidized 2-butene, 54% of the oxidized substrate was converted to
the epoxide, while the remaining 46% was converted to 2-butene-1-ol
(15). Because toluene monooxygenases also can hydroxylate
methyl groups (17), it is possible that a similar reaction
occurred in our 2-butene oxidation assays, thereby limiting the amount
of epoxide generated.
As stated previously, the accumulation of epoxides was transient. The
disappearance of the epoxides could be the result of either biological
or chemical reactions. Previous experiments with MMO found that no
diepoxide was formed from butadiene, and the potential for
MMO-catalyzed hydrolysis of BME was not suggested (15). It
was suggested, however, that BME may be too large to fit into the
active site of MMO. Conversely, van Hylckama et al. (20)
found that both ethene epoxide and cis-DCE epoxide served as
substrates for MMO, although the products of the reaction were not
identified. Here, when toluene-induced and uninduced cultures of G4 or
ENVBF1 were incubated with BME, the uninduced cells did not oxidize
BME, whereas the induced cells did. Furthermore, the presence of
toluene and butadiene inhibited the degradation of BME. These results
show that although BME may undergo slow chemical hydrolysis in the
presence of water at a neutral pH, toluene monooxygenase-expressing cells catalyze a more rapid oxidation of BME. Thus, it is likely that
toluene monooxygenase acts as the catalyst. Epichlorohydrin and butane
epoxide were also degraded by toluene monooxygenase-expressing organisms. We were unable to determine if the transformation of the
epoxides led to the hydrolysis of the epoxide ring, forming the
corresponding diol, or if the oxygenases hydroxylated the compounds at
one of the acyclic carbons.
Because of the growing interest in enantiomerically pure feedstocks for
both industrial and pharmaceutical chemical syntheses, we examined the
enantiomeric selectivity of the epoxidation reactions catalyzed by the
various toluene monooxygenases with 1,3-butadiene and 1-pentene as
substrates. In some cases, the toluene monooxygenases catalyzed
epoxidation reactions with a high degree of enantiomeric selectivity
(Table 3). For example, when the T2MO of G4 oxidized 1-pentene, only
one enantiomer of pentene epoxide could be detected. Thus, strain G4
may be a candidate catalyst for producing pentene epoxide for use in
pharmaceutical syntheses.
In this study, we examined variants of toluene monooxygenase that
oxidize the aromatic ring of toluene at all three possible positions
with three representatives, KR1, ENVPC5, and ENVBF1, of the
T4MO variety. (In a previous report [13], we suggested that the toluene monooxygenase of ENVBF1 was a T2MO, based on oxygen consumption studies following growth on toluene, but we have
since discovered that the cloned ENVBF1 toluene oxygenase genes
produce p-cresol during toluene oxidation.) When
1,3-butadiene served as the substrate, the three toluene monooxygenase
isoforms that oxidize chloroform (13) had the lowest degree
of enantiomeric selectivity. The enantiomeric selectivities were
similar to that of MMO [36% (R) and 64% (S)],
which also oxidizes chloroform (9). Therefore, it appears
that the enzyme plasticity that allows these related enzymes to oxidize
a wide range of substrates, including chloroform, may limit their
applicability for producing highly pure epoxide enantiomers.
With the exception of the oxidations catalyzed by the T2MO of G4, the
enantiomeric selectivities of the oxidations catalyzed by the wild-type
toluene monooxygenases are probably too low for them to be useful for
the commercial production of enantiomerically pure epoxides. However,
it might be possible to improve the enantiomeric selectivity of these
reactions through site-directed mutagenesis of toluene monooxygenases.
More research, including an evaluation of product yields, stability,
and toxicity, is needed to further understand the potential of the
toluene monooxygenases as biocatalysts for the production of
enantiomerically pure epoxides.
| |
ACKNOWLEDGMENTS |
This work was supported in part by an NSF Small Business
Innovative Research grant (DMI-9460076) to R.J.S. and an NSF Early Career grant (MCB-9733374) to B.G.F.
We thank G. Zylstra for kindly providing P. putida strains
F1 and strain GZ-9.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: 4100 Quakerbridge Rd., Lawrenceville, NJ 08648. Phone: (609) 936-9300. Fax:
(609) 936-9221. E-mail: Steffan{at}envirogen.com.
| |
REFERENCES |
| 1.
|
Archelas, A., and R. Furstoss.
1997.
Synthesis of enantiopure epoxides through biocatalytic approaches.
Annu. Rev. Microbiol.
51:491-525[CrossRef][Medline].
|
| 2.
|
Besse, P., and H. Veschambre.
1994.
Chemical and biological synthesis of chiral epoxides.
Tetrahedron
50:8885-8927[CrossRef].
|
| 3.
|
Byrne, A. M.,
J. J. Kukor, and R. H. Olsen.
1995.
Sequence analysis of the gene cluster encoding toluene-3-monooxygenase from Pseudomonas pickettii PKO1.
Gene
154:65-70[CrossRef][Medline].
|
| 4.
|
Fox, B. G.,
J. G. Borneman,
L. P. Wackett, and J. D. Lipscomb.
1990.
Haloalkene oxidation by the soluble methane monooxygenase from Methylosinus trichosporium OB3b: mechanistic and environmental implications.
Biochemistry
29:6419-6427[CrossRef][Medline].
|
| 5.
|
Gray, K. M., and E. P. Greenberg.
1992.
Physical and functional maps of the luminescence gene cluster in an autoinducer-deficient Vibrio fischeri strain isolated from a squid light organ.
J. Bacteriol.
174:4384-4390[Abstract/Free Full Text].
|
| 6.
|
Habets-Crutzen, A. Q. H.,
S. J. N. Carlier,
J. A. M. de Bont,
D. Wistuba,
V. Schurig,
S. Hartmans, and J. Tramper.
1985.
Stereospecific formation of 1,2-epoxypropane, 1,2-epoxybutane, and 1-chloro-2,3-epoxypropane by alkene utilizing bacteria.
Enzyme Microb. Technol.
7:17-21.
|
| 7.
|
Habets-Crutzen, A. Q. H.,
L. E. S. Brink,
C. G. van Ginkel,
J. A. M. de Bont, and J. Tramper.
1984.
Production of epoxides from gaseous alkenes by resting-cell suspensions and immobilized cells of alkene-utilizing bacteria.
Appl. Microbiol. Biotechnol.
20:245-250.
|
| 8.
|
Hareland, W. A.,
R. L. Crawford,
P. J. Chapman, and S. Dagley.
1975.
Metabolic function and properties of 4-hydroxyphenylacetic acid 1-hydroxylase from Pseudomonas acidovorans.
J. Bacteriol.
121:272-285[Abstract/Free Full Text].
|
| 9.
|
Jahng, D., and T. K. Wood.
1994.
Trichloroethylene and chloroform degradation by a recombinant pseudomonad expressing soluble methane monooxygenase from Methylosinus trichosporium OB3b.
Appl. Environ. Microbiol.
60:2473-2482[Abstract/Free Full Text].
|
| 10.
|
Lalonde, J.
1997.
Enzyme catalysis: cleaner, safer, energy efficient.
Chem. Eng.
9:108-112.
|
| 11.
|
Lange, C. C., and L. P. Wackett.
1997.
Oxidation of aliphatic olefins by toluene dioxygenase: enzyme rates and product identification.
J. Bacteriol.
179:3858-3865[Abstract/Free Full Text].
|
| 12.
|
May, S. W.,
M. S. Steltenkamp,
R. D. Schwartz, and C. J. McCoy.
1976.
Stereoselective formation of diepoxides by an enzyme system of Pseudomonas olevorans.
J. Am. Chem. Soc.
98:7856-7858[CrossRef][Medline].
|
| 13.
|
McClay, K.,
B. G. Fox, and R. J. Steffan.
1996.
Chloroform mineralization by toluene-oxidizing bacteria.
Appl. Environ. Microbiol.
62:2716-2722[Abstract].
|
| 14.
|
Oldenhuis, R.,
J. Y. Oedzes,
J. J. van der Waarde, and D. B. Janssen.
1991.
Kinetics of chlorinated hydrocarbon degradation by Methylosinus trichosporium OB3b and toxicity of trichloroethylene.
Appl. Environ. Microbiol.
57:7-14[Abstract/Free Full Text].
|
| 15.
|
Ono, M., and I. Okura.
1990.
On the reaction mechanisms of alkene epoxidation with Methylosinus trichosporium OB3b.
J. Mol. Catal.
61:113-122[CrossRef].
|
| 16.
|
Panke, S.,
B. Witholt,
A. Schmid, and M. G. Wubbolts.
1998.
Towards a biocatalyst for (S)-styrene oxide production: characterization of the styrene degradation pathway of Pseudomonas sp. strain VLB120 Appl.
Environ. Microbiol.
64:2032-2043.
|
| 17.
|
Pikus, J. D.,
J. M. Studts,
K. McClay,
R. J. Steffan, and B. G. Fox.
1997.
Changes in the regiospecificity of aromatic hydroxylation produced by active site engineering in the diiron enzyme toluene 4-monooxygenase.
Biochemistry
36:9283-9289[CrossRef][Medline].
|
| 18.
|
Shields, M. S.,
S. O. Montgomery,
P. J. Chapman,
S. M. Cuskey, and P. H. Pritchard.
1989.
Novel pathway of toluene catabolism in the trichloroethylene-degrading bacterium G4.
Appl. Environ. Microbiol.
55:1624-1629[Abstract/Free Full Text].
|
| 19.
|
Stinson, S. C.
1998.
Counting on chiral drugs.
Chem. Eng. News
76:1-36.
|
| 20.
|
van Hylckama,
J. E. T. Vlieg,
W. de Koning, and D. B. Janssen.
1996.
Transformation kinetics of chlorinated ethenes by Methylosinus trichosporium OB3b and detection of unstable epoxides by on-line chromatography.
Appl. Environ. Microbiol.
62:3304-3312[Abstract].
|
| 21.
|
Yeager, C. M.,
P. J. Bottomley,
D. J. Arp, and M. R. Hyman.
1999.
Inactivation of toluene 2-monooxygenase in Burkholderia cepacia G4 by alkynes.
Appl. Environ. Microbiol.
65:632-639[Abstract/Free Full Text].
|
| 22.
|
Yen, K.-M.,
M. R. Karl,
L. M. Blatt,
M. J. Simon,
R. B. Winter,
P. R. Fausset,
H. S. Lu,
A. A. Harcourt, and K. K. Chen.
1991.
Cloning and characterization of a Pseudomonas mendocina KR1 gene cluster encoding toluene-4-monooxygenase.
J. Bacteriol.
173:5315-5327[Abstract/Free Full Text].
|
| 23.
|
Zhou, N.-Y.,
A. Jenkins,
K. N. Chan,
C. K. Chion, and D. J. Leak.
1998.
The alkene monooxygenase from Xanthobacter PY2 is a binuclear non-haem iron protein closely related to toluene-4-monooxygenase.
FEBS Lett.
430:181-185[CrossRef][Medline].
|
| 24.
|
Zylstra, G. J.,
W. R. McCombie,
D. T. Gibson, and B. A. Finette.
1988.
Toluene degradation by Pseudomonas putida F1: genetic organization of the tod operon.
Appl. Environ. Microbiol.
54:1498-1503[Abstract/Free Full Text].
|
Applied and Environmental Microbiology, May 2000, p. 1877-1882, Vol. 66, No. 5
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Abstract
Introduction
