INTRODUCTION

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The aerobic bacterial degradation of nonactivated aromatic compounds is usually initiated by dioxygenases that incorporate two hydroxyl groups into the aromatic substrate; the products of such reactions are chiral cis-dihydrodihydroxy derivatives, which also are called cis-dihydrodiols (10). The oxidation of aromatic compounds to cis-dihydrodiols is of special interest for biotechnological as well as chemical applications, because single cis-dihydrodiols have been shown to be valuable building blocks for stereoselective synthesis of biologically active molecules containing multiple chiral centers (6, 8, 12, 16, 18, 22).

Several multicomponent dioxygenases that dihydroxylate aromatic compounds have been described (7). Toluene dioxygenases (TDO), naphthalene dioxygenase (NDO), and biphenyl dioxygenase (BDO) are the best known members of this class of dioxygenases (12). The components of these oxygenase complexes have been purified, and the encoding genes have been cloned and expressed in Escherichia coli (12). Experiments on substrate specificity showed that TDO from Pseudomonas putida F39/D and P. putida UV4 oxidize monosubstituted benzenes, except fluorobenzene, to 3-substituted cis-benzene dihydrodiols with an S configuration at the C-1 atom (3, 4, 15). BDO from Pseudomonas sp. strain LB400 dihydroxylates and dechlorinates chlorinated biphenyls (11) to the respective 2,3 or 3,4 cis-diols, whereby dihydroxylation and dechlorination happen on both phenyl rings. NDO from Pseudomonas sp. strain NCIB 9816 has a relaxed substrate specificity and catalyzes the dioxygenation of naphthalene to (+)-cis-(1R,2S)dihydroxy-1,2-dihydronaphthalene. Many related 2- and 3-ring aromatic and hydroaromatic compounds are also substrates and are turned over to the respective cis-diols (14). A mutant of the not-so-well-characterized Pseudomonas fluorescens N3 was used in the bioconversion of several naphthalene derivatives to the corresponding cis-dihydrodiols on a milligrams-to-grams scale (2). A recent review lists more than 140 diols that have been described to date (12). Although only a few of them have been used as synthons, the further development of the area depends on the discovery of novel dioxygenase reactions and on the development of biotechnological processes to produce the different metabolites.

Werlen et al. recently described the cloning and expression in E. coli DH5(pTCB144) of the chlorobenzene dioxygenase (CDO) of Pseudomonas sp. strain P51 (21). Analysis of the genes showed that CDO is a three-component aromatic ring dioxygenase. It consists of the gene products of tcbAa, encoding the large subunit of the terminal oxygenase; tcbAb, encoding the small subunit; tcbAc, encoding the ferredoxin; and tcbAd, encoding the NADH reductase. Homology comparisons indicate that these genes and gene products are most closely related to those of TDO of P. putida F1 (todC1C2BA) and are distantly related to those of NDO and BDO (21). Here we report the ability of the CDO system of the recombinant E. coli DH5(pTCB144) to oxidize different classes of aromatic compounds to the corresponding cis-dihydrodiols. We characterized each of the 48 reaction products as thoroughly as possible. The CDO system turned out to be well suited for the production of dihydrodiols, and further work to scale up production is under way.

	MATERIALS AND METHODS

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Microorganisms, growth conditions, and biotransformation procedure. Single colonies of E. coli DH5(pTCB144) were taken from agar plates and transferred to Erlenmeyer flasks containing 250 ml of sterile Luria-Bertani medium (17) with 50 mg of ampicillin/liter. Cultures were incubated at 25°C on a shaker (110 rpm) for 36 h, by which time the optical density at 578 nm (OD₅₇₈) had reached levels between 3.5 and 4.0. The cultures were then centrifuged at 20,000 × g for 12 min. The supernatant was removed, and cells were washed with M9 mineral salts medium (17). After the wash, the cells were resuspended in M9 medium supplemented with 1 mM glucose, and the OD₅₇₈ was adjusted to approximately 1.0. These cells were used for biotransformation incubations as follows. One hundred milliliters of the washed suspension was transferred to 500-ml Erlenmeyer flasks. Aromatic substrates were added from stock solutions in methanol (35 mM) prior to bacterial suspension. Methanol was allowed to evaporate. Final substrate concentrations were 0.35 mM. Experiments for determination of product-building rates were performed with 1.0 mM concentrations of the substrate. Samples were taken approximately every 45 min and centrifuged. The supernatants were frozen until needed for analysis. Biphenyl, which was known to be transformed by CDO (21), was used as a positive control for all experiments. Experiments were carried out at 25°C.

Analytical procedures. (i) Rate determination. The formation of metabolites was measured by high-pressure liquid chromatography (HPLC). Samples of the supernatants from the biotransformation incubations (2 ml) were filtered through 0.20-µm-pore-size membrane filters (FP 030/3; Schleicher & Schuell, Dassel, Germany) or centrifuged at 25,000 × g for 2 min and analyzed on a Waters 625 LC HPLC system equipped with a WISP 700 autosampler and a 901 photodiode array detector (Waters Millipore Corp., Milford, Mass.). Separation was done on a C₁₈ reversed-phase column (Macherey-Nagel, Düren, Germany) as described elsewhere (21). In order to determine product-building rates, we quantified the transformation products of benzene, fluorobenzene, and naphthalene with the help of authentic standards. The transformation products of toluene and of ethyl-, chloro-, and bromobenzene were acidified, and the resulting phenols were analyzed and quantified accordingly.

(ii) GC-MS. For gas chromatography-mass spectrometry (GC-MS) analysis, cis-dihydrodiols were extracted from the supernatant of the incubation mixture with an equal volume of ethyl acetate. The ethyl acetate extract was dried with sodium sulfate and evaporated to dryness under a gentle stream of nitrogen at 40°C. The residue was dissolved in 100 µl of N,N-dimethylformamide (DMF). One hundred microliters of a solution of recrystallized n-butylboronic acid (approximately 500 µg of n-butylboronic acid/ml dissolved in DMF) was added, and the mixture was heated to 70°C for 15 min to form the n-butylboronate derivatives (BB derivatives). The free hydroxy groups of the BB derivatives of the transformation products of the monohydroxybiphenyls, phenol, cresols, and benzyl alcohol were then derivatized with N,O-bis(trimethylsilyl)-trifluoroacetamide (TMS) as described elsewhere (13). Samples of the BB derivatives were diluted with cyclohexane (at least 15-fold), 0.5 µl of the diluted samples was injected at 50°C onto a PS09 fused silica capillary column (length, 15m; inner diameter, 0.25 mm; film thickness, 0.25 µm), and the column temperature was increased to 250°C at 10°C/min. Mass spectra were obtained with an ITD 800 (ion trap detection) mass spectrometer (Finnigan, MAT, San Jose, Calif.) coupled with an HRGC 5160 Mega Series gas chromatograph (Carlo Erba Instruments, Milan, Italy).

(iii) ¹H NMR spectroscopy. Nuclear magnetic resonance (NMR) spectra were recorded on a Bruker ASX 400-MHz NMR spectrometer at ambient temperature. The solvents were methanol-d₄ or acetonitrile-d₃. Chemical shifts are given in parts per million relative to tetramethylsilane (at 0 ppm).

(iv) Protein determination. Protein contents were determined by the method of Bradford (5) with bovine serum albumin as the standard.

(v) Optical rotations. Optical rotations were measured in methanol solutions with a Polarimeter 241 (Perkin-Elmer International Inc., Rotkreuz, Switzerland).

Isolation of products. In order to isolate some of the transformation products in the milligram range, the supernatants of the incubation mixtures (1.5 to 2.5 liters) were extracted twice with 1/2 volume of ethyl acetate. Prior to the extraction, a sufficient amount of Na₂SO₄ was added in order to improve the transfer of the cis-dihydrodiols into the organic phase and to prevent the formation of emulsions. The ethyl acetate was evaporated in a rotary evaporator to dryness, and the remains were redissolved in methanol and filtered (FP 030/3; pore size, 0.2 µm; Schleicher & Schuell). Samples of the filtered methanol extracts (500 µl) were injected on a semipreparative C₁₈ column (ET 250/21, Nucleosil 100-7; Macherey-Nagel). Compounds were eluted by running a linear gradient from 70% (vol/vol) water and 30% methanol to 30% water and 70% methanol in 40 min.

Chemicals. Monochlorobiphenyls and 4,4'-dichlorobiphenyl were obtained from Johnson & Matthey (Karlsruhe, Germany); 2,2'-dichlorobiphenyl and 2,2',6-trichlorobiphenyl were obtained from Promochem (Wesel, Germany); 4-chlorotoluene, 2,2-bis(4-chlorophenyl)-1,1-dichlorethane (DDE), pyrene, 1,1-bis(4-chlorophenyl)-2,2,2-trichlorethane (DDT), 4-biphenylcarbonitrile, all dihydroxybiphenyls, 1,4-dichlorobenzene, 1,2,3-trichlorobenzene, dibenzofuran (DBF), diphenylether, cis-1,2-dihydrocatechol, 2,4-dichlorophenoxyacetic acid, and 2-hydroxydiphenylmethane were obtained from Aldrich (Buchs, Switzerland); 4-fluoroaniline, dibenzo-p-dioxin, and 2,6-dichlorobiphenyl were obtained from Socochim (Lausanne, Switzerland); 4-bromo-2-methylphenol, all tetrachlorobenzenes, 2,3-dichlorobiphenyl, and 2,4,5-trichlorobiphenyl were obtained from Riedel de Häen (Seelze, Germany); 2-bromo-4-methylphenol was obtained from Janssen Chimica (Geel, Belgium). 3-Chlorodibenzofuran was a gift from Hauke Harms (Swiss Federal Institute for Environmental Sciences and Technology [EAWAG]). Standards of (±)-cis-1,2-dihydroxy-1,2-dihydronaphthalene, (±)-cis-1,2-dihydroxy-1,2,3,4-tetrahydronaphthalene, (+)-cis-(2R,3S)-dihydroxy-2,3-dihydrobiphenyl, and ()-cis-(2S,3R)-dihydroxy-2,3-dihydrobiphenyl were gifts from S. M. Resnick and D. T. Gibson. All other chemicals were obtained from Fluka (Buchs, Switzerland).

	RESULTS

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Identification of reaction products. Dihydrodiol metabolites were identified by GC-MS of their respective BB derivatives and, when possible, by ¹H NMR spectroscopy. BB derivatives of cis-dihydrodiol isomers were separated on a GC column with an achiral stationary phase and could be distinguished from those of trans-dihydrodiols because of significant mass differences between their molecular ions (19).

View larger version (10K): [in this window] [in a new window]   FIG. 1.   Designation of the protons of mono- and 1,4-disubstituted cis-benzene dihydrodiols, formed by CDO.

Transformation of substituted biphenyls. CDO dihydroxylated biphenyl and several monosubstituted biphenyls (Table 1). Biphenyl was oxidized to enantiomerically pure (+)-cis-(2R,3S)-dihydroxy-2,3-dihydrobiphenyl. Oxidation of monochlorobiphenyls occurred on the unsubstituted ring at the 2,3 position. Measurements of the optical rotations of the monochlorinated cis-biphenyl dihydrodiol products confirmed that these products were nonracemic (Table 1). Several other monosubstituted biphenyls were oxidized by CDO. Higher substituted biphenyls (2,3- 2,6-, 2,2'-, and 4,4'-dichlorobiphenyl; 2,2',6- and 2,4,5-trichlorobiphenyl; 2,2'-, 4,4'-, and 2,5- dihydroxybiphenyl) were not transformed by CDO. Neither dihydroxylation nor dechlorination products were detected by GC-MS analysis.

View this table: [in this window] [in a new window]   TABLE 1.   Products formed from substituted biphenyls by whole cells of E. coli DH5(pTCB 144), which contains the CDO of Pseudomonas sp. strain P51

Transformation of substituted benzenes. CDO oxidized benzene, 1,4-dichlorobenzene, fluorobenzene, 1-chloro-4-iodobenzene, 1-bromo-4-iodobenzene, bromobenzene, styrene, anisol, and ethylbenzene to cis-dihydrodiol derivatives (data not shown). Most substrates were dihydroxylated to one cis-dihydrodiol isomer. As these products were previously described (12), they were not further investigated. Table 2 shows the mass spectral properties of the BB derivatives of interesting new cis-dihydrodiols. The data show that CDO is able to oxidize tri- and tetrachlorobenzene, benzonitrile, benzyl chloride, benzyl cyanide, and diphenylmethane to cis-dihydrodiols. CDO oxidized monosubstituted benzenes faster than benzene (Table 3). Compounds with polar substituents, such as phenol, 2,4-dichlorophenoxyacetic acid, benzyl alcohol, aniline, and 4-fluoroaniline did not serve as substrates for CDO. CDO did not oxidize 1,2,3,5-tetrachlorobenzene, 2-hydroxydiphenylmethane, DDE, or DDT.

View this table: [in this window] [in a new window]   TABLE 2.   Products formed from substituted benzenes by whole cells of E. coli DH 5(pTCB 144), which contains the CDO of Pseudomonas sp. strain P51

Transformation of substituted toluenes. Many substituted toluenes served as substrates for CDO. Incubation of toluene, 4-fluorotoluene, 4-bromotoluene, and 4-iodotoluene produced one single cis-dihydrodiol isomer [(+)-cis-(2R,3S)-dihydroxy-I-methylcyclohexa-4,6-diene and cis-4-bromo-2,3-dihydroxy-1-methylcyclohexa-4,6-diene for toluene and 4-bromotoluene, respectively], whereas incubation of 4-chlorotoluene produced all three possible isomers. E. coli DH5(pTCB144) oxidized para-substituted toluenes to the same major and minor enantiomers as P. putida F39/D, whereby the EE values of the products formed by E. coli DH5(pTCB144) were usually higher (Table 4). The separation by GC of the BB derivatives of cis-4-bromo-2,3-dihydroxy-1-methyl-cyclohexa-4,6-diene, which was formed by oxidation of 4-bromotoluene, is shown in Fig. 2.

View larger version (10K): [in this window] [in a new window]   FIG. 2.   Determination of the enantiomeric composition of cis-4-bromo-2,3-dihydroxy-1-methyl-cyclohexa-4,6-diene, which is formed by oxidation of 4-bromotoluene by CDO. Enantiomers were separated as their BB derivatives. The mass spectra of the enantiomers were identical. The major enantiomer was formed with an EE of 77%, which was calculated by integration of the peak areas of the major and minor enantiomer. The isomeric structure of the product was determined by 1H NMR analysis.

Transformation of additional aromatic substrates. CDO oxidized several additional aromatic substrates to cis-dihydrodiols. Naphthalene was oxidized to enantiomerically pure (+)-cis-(1R,2S)-dihydroxy-1,2-dihydronaphthalene (Fig. 3). It was formed at a rate of 22.8 nmol/min · mg of protein¹, which is much higher than the transformation rates of benzene and several monosubstituted benzenes (Table 3).

View larger version (15K): [in this window] [in a new window]   FIG. 3.   GC-MS analysis and determination of the absolute configuration of cis-1,2-dihydroxy-1,2-dihydronaphthalene. (A) Separation of the BB derivatives of an authentic standard of (±)-cis-1,2-dihydroxy-1,2-dihydronaphthalene. The mass spectra of the two compounds were identical. (B) BB derivative of an authentic standard of (+)-cis-(1R,2S)-dihydroxy-1,2-dihydronaphthalene. (C) BB derivative of the oxidation product formed from naphthalene by CDO. The BB derivatives in panels B and C had identical retention times and identical mass spectra.

View this table: [in this window] [in a new window]   TABLE 5.   Products formed from various aromatic substrates by whole cells of E. coli DH 5(pTCB144), which contains the CDO of Pseudomonas sp. strain P51

	DISCUSSION

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We showed that CDO from Pseudomonas sp. strain P 51, which was cloned into E. coli DH5(pTCB144) (21), was able to oxidize various substituted aromatic compounds to cis-dihydrodiols. The absolute configurations of several products were determined. They were identical to those of the products formed by TDO and NDO (1, 3, 15, 20, 24, 25). NMR spectra were recorded if enough product (ca. 1 mg) was available. The J-coupling pattern among the olefinic protons was used to assign the positions of the hydroxy groups. For some compounds (e.g. chlorobiphenyls and bromobenzene) most of the long-range J couplings were resolved as well. For products such as dihydrodihydroxytoluene or dihydrodihydroxyfluorobenzene, long-range J couplings manifested themselves in 1H,1H-cosy correlation signals but could not be extracted from the one-dimensional NMR spectra directly.

We were able to separate cis-dihydrodiol enantiomers by GC. Advantages of the method were the low detection limit (<0.5 µg of the derivative), the fact that no exhaustive isolation and cleanup was needed, and the fact that derivatization could be done with ordinary laboratory equipment. Several standards of the BB derivatives of racemic or nonracemic mixtures of cis-dihydrodiol enantiomers [e.g., (±)-cis-1,2-dihydroxy-1,2-dihydronaphthalene (Fig. 3), (±)-cis-1,2-dihydroxy-1,2,3,4-tetrahydronaphthalene, (±)-cis-2,3-dihydroxy-2,3-dihydrobiphenyl, and (±)-cis-1,2-indandiol] were tested and could be successfully separated. Even high EE values were quantified. For example, the EE values of the cis-dihydrodiol products of fluorobenzene and 4-iodotoluene turned out to be 95 and 98%, respectively. The analyses were reproducible with a relative standard deviation of 3%.

One disadvantage of the method was that a single peak in a gas chromatogram does not automatically prove enantiomeric purity of the measured compound. To this end, standards of both enantiomers would be needed to ensure that in principle they can be separated. Many measurements gave single peaks, e.g., the BB derivatives of cis-chlorobenzene dihydrodiols or of the monosubstituted cis-biphenyl dihydrodiols. Since authentic standards of both enantiomers of these products were unavailable, we could not unequivocally prove enantiopurity. So far, all available standards were separated and quantifications of even high EE values were reproducible. Therefore, it is quite likely that most cis-dihydrodiols that eluted as single peaks also are pure enantiomers.

CDO oxidized various 1,4-disubstituted benzenes to two cis-dihydrodiol enantiomers. With the exception of the oxidation product of 4-fluorotoluene, the products that were formed by CDO had higher EE values than those formed by TDO of P. putida UV4 and P. putida F39/D (Table 4). Boyd et al. (4) concluded that the EE of 1,4-disubstituted cis-benzene dihydrodiols formed by TDO of P. putida UV4 is largely controlled by steric effects. Obviously, there have to be additional parameters that influence the EE of 1,4-disubstituted cis-benzene dihydrodiols formed by CDO.

The formation rates of the transformation products of benzene, several monosubstituted benzenes, and naphthalene are given in Table 3. As indicated in an earlier study (21), naphthalene is turned over faster by CDO than benzene and monosubstituted benzenes. The formation rates of the monosubstituted cis-benzene dihydrodiols are higher than that of cis-benzene dihydrodiol. We assume that the large substituent helps the substrate to fit into the active binding site of the enzyme. However, the differences in the transformation rates between the monosubstituted benzenes cannot be exclusively explained by varying sizes of the substituent. We postulate that the rate of formation of monohalogenated cis-benzene dihydrodiols by CDO increases with smaller size and with lower electronegativity of the halogen substituent. Both influences compensate for each other in the case of a chlorine or fluorine substituent.