Trichloroethylene Oxidation by Purified Toluene 2-Monooxygenase ...

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Chloral hydrate and dichloroacetic acid were not detected. This finding differs from that with soluble methane monooxygenase and cytochrome P-450.
JOURNAL OF BACTERIOLOGY, Jan. 1997, p. 90–96 0021-9193/97/$04.0010 Copyright q 1997, American Society for Microbiology

Vol. 179, No. 1

Trichloroethylene Oxidation by Purified Toluene 2-Monooxygenase: Products, Kinetics, and Turnover-Dependent Inactivation LISA M. NEWMAN†

AND

LAWRENCE P. WACKETT*

Departments of Biochemistry and Microbiology, the Biological Process Technology Institute, and the Center for Biodegradation Research and Informatics, University of Minnesota, St. Paul, Minnesota 55108 Received 23 July 1996/Accepted 21 October 1996

Trichloroethylene is oxidized by several types of nonspecific bacterial oxygenases. Toluene 2-monooxygenase from Burkholderia cepacia G4 is implicated in trichloroethylene oxidation and is uniquely suggested to be resistant to turnover-dependent inactivation in vivo. In this work, the oxidation of trichloroethylene was studied with purified toluene 2-monooxygenase. All three purified toluene 2-monooxygenase protein components and NADH were required to reconstitute full trichloroethylene oxidation activity in vitro. The apparent Km and Vmax were 12 mM and 37 nmol per min per mg of hydroxylase component, respectively. Ten percent of the full activity was obtained when the small-molecular-weight enzyme component was omitted. The stable oxidation products, accounting for 84% of the trichloroethylene oxidized, were carbon monoxide, formic acid, glyoxylic acid, and covalently modified oxygenase proteins that constituted 12% of the reacted [14C]trichloroethylene. The stable oxidation products may all derive from the unstable intermediate trichloroethylene epoxide that was trapped by reaction with 4-(p-nitrobenzyl)pyridine. Chloral hydrate and dichloroacetic acid were not detected. This finding differs from that with soluble methane monooxygenase and cytochrome P-450 monooxygenase, which produce chloral hydrate. Trichloroethylene-dependent inactivation of toluene 2-monooxygenase activity was observed. All of the protein components were covalently modified during the oxidation of trichloroethylene. The addition of cysteine to reaction mixtures partially protected the enzyme system against inactivation, most notably protecting the NADH-oxidoreductase component. This suggested the participation of diffusible intermediates in the inactivation of the oxidoreductase. ene dioxygenase oxidizes TCE to formic and glyoxylic acids in a 2:1 ratio (20). In both cases, turnover-dependent enzyme inactivation occurs in vitro (12, 20) and in vivo (29, 37). It has been suggested that enzyme inactivation results largely from reactive acyl chloride intermediate compounds that alkylate the proteins. These observations suggest limits to the practicality of bioremediating TCE via bacteria containing nonspecific catabolic oxygenases. Against this backdrop, Burkholderia cepacia G4 oxidizes TCE, purportedly via toluene 2-monooxygenase, with little apparent inactivation (11, 32). This has stimulated the use of that bacterium in microcosoms and bioreactors developed specifically for treating TCE-contaminated water (10, 18, 19, 40). The promulgation of in vivo TCE remediation methods based on the catalytic activity of toluene 2-monooxygenase is occurring in the absence of information about the enzymatic reaction products and related kinetic data. The present study is designed to fill that gap. The three components of toluene 2-monooxygenase were recently purified, and its structure and catalytic properties with toluene were reported (28). In vitro reconstitution of efficient toluene oxidation activity requires (i) a flavo-iron-sulfur protein, (ii) a small nonchromophoric protein, and (iii) an a2b2g2 binuclear-iron protein. In the present study, the reaction of TCE with purified toluene 2-monooxygenase components was investigated with respect to (i) steady-state kinetic parameters, (ii) the reaction products, and (iii) enzyme inactivation that occurs during in vitro TCE oxidation.

Trichloroethylene (TCE) is a widely used industrial solvent that has become a common contaminant of drinking water. The mammalian toxicity of TCE is low, and epidemiological evidence suggests it is not a human carcinogen (14). Of most concern is the observation that anaerobic bacteria in groundwater sometimes catalyze reductive dechlorination of TCE, yielding vinyl chloride (35). The latter compound is a potent human carcinogen (23, 30). This gives impetus for extensive efforts to remediate TCE-contaminated environments, and some of the efforts use bacteria with enzymes capable of oxidizing TCE. There are no well-documented reports of bacterial growth on TCE as the sole carbon and energy source, but some organisms fortuitously oxidize the compound via nonspecific catabolic oxygenases (5–7, 9). For example, soluble methane monooxygenase and toluene dioxygenase each oxidize dozens, if not hundreds, of different organic compounds, including TCE (13, 17, 21, 38). This has focused efforts on using organisms that express these enzymes for TCE bioremediation. Soluble methane monooxygenase is a three-protein enzyme system containing a binuclear-iron hydroxylase component. Toluene dioxygenase is also a three-protein enzyme system, thought to contain a dihydroxylase component with a mononuclear iron center. Purified soluble methane monooxygenase oxidizes TCE at a rate of 70% the Vmax of methane with the production of TCE epoxide and 2,2,2-trichloroacetaldehyde (chloral) (12). Tolu* Corresponding author. Mailing address: Department of Biochemistry and Biological Process Technology Institute, University of Minnesota, 240 Gortner Laboratory, 1479 Gortner Ave., St. Paul, MN 55108. Phone: (612) 625-3785. Fax: (612) 625-1700. E-mail: wackett @biosci.cbs.umn.edu. † Present address: Center for Agricultural Molecular Biology, Cook College, Rutgers University, New Brunswick, NJ 08903.

MATERIALS AND METHODS Growth of bacterial strains. B. cepacia G4 was provided by Malcolm Shields (University of West Florida). B. cepacia G4 was maintained on minimal salts buffer (MSB) (33) agar plates containing 20 mM lactate. For liquid growth of B. cepacia G4, 50 ml of MSB medium in a 250-ml flask containing 20 mM lactate was inoculated and incubated, with shaking at 200 rpm, for 24 h at 238C. For

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VOL. 179, 1997 growth on toluene, flasks containing 250 to 500 ml of MSB medium were inoculated with this overnight culture and incubated under the same conditions with toluene as the sole carbon source, supplied as a vapor by suspension of a vapor bulb over the liquid phase of the culture. The liquid phase did not exceed 25% of the volume of the flask. Cells were harvested by centrifugation at 5,000 3 g for 15 min after the cells had attained an optical density at 600 nm of 2.0. The cell pellet was washed with 25 mM MOPS (morpholinepropanesulfonic acid [pH 7.5]) and resuspended in 25 mM MOPS at pH 7.5 to an optical density at 600 nm of 10. Cultures were assayed for expression of toluene 2-monooxygenase activity by a rapid colorimetric assay which detects the enzymatic formation of naphthols from naphthalene (3). Aliquots of cultures were incubated in test tubes with one or two crystals of naphthalene for 15 minutes, with shaking at 200 rpm, after which 100 ml of freshly hydrated 4 mM tetrazotized o-dianisidine was added. Immediate development of a purple color indicated the culture was actively expressing toluene 2-monooxygenase. Purification of enzyme components. Toluene 2-monooxygenase components were purified and analyzed for activity as described previously, except the pH of the buffer was 7.8 (28). In these studies, the specific activity of the toluene 2-monooxygenase with o-cresol was 202 nmol/min mg of hydroxylase21, and the iron content was 7.2 nmol of iron/nmol of hydroxylase. This was somewhat higher in iron content and specific activity than previously reported (28). Steady-state kinetics parameters of TCE oxidation. Steady-state kinetic parameters were determined with purified toluene 2-monooxygenase enzyme components. Reaction mixtures contained, in a 200-ml volume of 100 mM MOPS at pH 7.0, 0.5 mM hydroxylase, 2.0 mM small component, 2.7 mM reductase, 2.5 mM NADH, and various concentrations of [14C]TCE added from 17 and 100 mM stock solutions made up in N,N9-dimethylformamide (DMF). The percentage of DMF in all vials was #1.0%. Reactions were performed in 1.8-ml septum-sealed vials, initiated upon addition of NADH, and incubated with shaking at 200 rpm at 238C. After 5 min, aliquots of the reaction mixture were spotted onto 1-cm2 plastic-backed silica thin-layer chromatography sheets, and any unreacted substrate was allowed to volatilize for 25 min in a fume hood. The dry thin-layer chromatography squares were then placed in 7 ml of scintillation fluid and analyzed for radioactivity. Product quantitation required correction to account for the volatile products; the molar concentration of nonvolatile products was divided by 0.43. This number is based on the product ratios shown in Table 2. The dimensionless Henry’s Law constant for TCE, 0.40, was used to calculate the amount of TCE in the aqueous phase (11). Kinetic data were treated by the graphical methods of Lineweaver-Burk and Eisenthal and Cornish-Bowden with similar results (31). Substrate inhibition was observed at high TCE concentrations. The kinetic parameters were derived from rates obtained at TCE concentrations showing hyperbolic curves in plots of rate versus substrate concentration. The kinetic parameters for TCE were determined with fixed concentrations of NADH and oxygen and are therefore apparent values. TCE oxidation products. Whole-cell assays were performed with 10-ml crimpsealed septum vials to which 2.0 ml of washed uninduced or toluene-induced cell suspensions was added (3, 36). [14C]TCE was added from a 10 mM stock solution in DMF to a final concentration of 100 mM. Vials containing viable and heatkilled controls were incubated, with shaking at 200 rpm, for 0, 30, and 60 min at 238C. Incubation mixtures were analyzed for 14CO2 production, radiolabeled acidic products, and incorporation of radiolabel into cell material as described in “Analytical methods.” Reconstituted enzyme reaction mixtures, containing 170 mM radiolabeled TCE added from a 10 mM stock solution in DMF, were incubated at 238C with shaking at 200 rpm. The reaction mixtures contained, in a 400-ml final volume, 2 mM hydroxylase, 4 mM small component, and 5 mM reductase in 100 mM MOPS (pH 7.0). The reactions were initiated by the addition of 2.5 mM NADH. Control mixtures contained various combinations of the reaction components to determine the requirements for oxidation of TCE. At appropriate time intervals, the reaction mixtures were transferred to clean Eppendorf tubes containing 8 ml of 1 N H2SO4 and analyzed by high-pressure liquid chromatography (HPLC) as described in “Analytical methods.” A nonradiolabeled reaction mixture was used to monitor the time course of the reaction by headspace gas chromatography analysis. Radiolabeled CO2 was determined as described below. Oxidation of NADH was determined by observing the decrease in A340. In experiments to determine carbon monoxide production, nonradiolabeled TCE was added to a final concentration of 2 mM. Confirmation of the products formate and glyoxylate. Glyoxylate and formate were positively identified through the addition of glyoxylate reductase and formate dehydrogenase to aliquots of the [14C]TCE oxidation reaction mixture. To 100 ml of reaction mixture, 0.2 U of glyoxylate reductase or formate dehydrogenase was added with 1 mM NADH or NAD1, respectively. The aliquots were incubated for 1 h at 238C. The reaction mixture containing formate dehydrogenase was incubated in a septum-sealed vial under a constant stream of air, which was analyzed for radiolabeled CO2. The reaction mixture containing glyoxylate reductase was stopped after 1 h by the addition of 2 ml of 1 N H2SO4, and precipitated protein was removed by centrifugation. Both reaction mixtures were analyzed by HPLC with an organic acid column as described in “Analytical methods.” Determination of TCE epoxide formation. TCE epoxide formation was detected with 4-(p-nitrobenzyl)pyridine by the method of Miller and Guengerich

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(24). The reaction mixtures contained, in a 200-ml final volume, 8.0 mM hydroxylase, 16 mM small component, and 16 mM reductase, in 100 mM MOPS (pH 7.0). TCE was added in DMF to a final concentration of 2 mM. The reactions were initiated by the addition of 2.5 mM NADH. The soluble methane monooxygenase enzyme system was used as a positive control for the formation of TCE epoxide, and reaction conditions were as previously reported (12). Control reactions consisted of the above reaction conditions without the addition of either NADH or enzyme. The concentration of the adduct formed with TCE epoxide was estimated with ε540 5 24 mM21 cm21 (8). Covalent labeling of enzyme components. Covalent labeling of the individual enzyme components was analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Gradient (5 to 30%) gels containing N,N9-diallyltartardiamide, in place of N,N9-methylene-bisacrylamide, were used (41). Aliquots of the reaction mixture were electrophoresed, and the gels were stained with Coomassie blue. The protein bands were excised and dissolved in 2% periodic acid, and the eluted protein was analyzed for radioactivity. Amounts of radiolabel associated with each protein band were normalized for the size of the protein by determining the number of counts per minute per surface area (As). As for each protein component was determined by the formula As (Å2) 5 6.3(Mw)0.73 (4). Time course of enzyme inactivation. Possible TCE-dependent inactivation of the enzyme components of toluene 2-monooxygenase was analyzed with protocols adapted from those previously described (12). Reaction mixtures contained 8.0 mM hydroxylase, 16 mM small component, 16 mM reductase, and 3.5 mM NADH, in 100 mM MOPS (pH 7.0). All reaction mixtures were incubated at 238C, with or without the addition of 2 mM TCE. Superoxide dismutase (100 U/ml) and catalase (100 U/ml) were added to all reaction vials to prevent organic substrate-independent enzyme inactivation. Triplicate reactions were assayed, and reactions were initiated by the addition of NADH. At appropriate time intervals, aliquots were withdrawn and assayed for the ability to oxidize o-cresol to 3-methylcatechol with a coupled enzyme assay (28). The coupled enzyme assay was performed as described previously (28), except the pH of the buffer used was 7.8. Oxidation of TCE was followed by headspace gas chromatography analysis as described in “Analytical methods.” The addition of 10 mM L-cysteine was analyzed for the ability to prevent enzyme inactivation. To determine the activity remaining for each enzyme component, aliquots of the inactivation reaction mixtures described above were withdrawn at 30 min and assayed in the following manner. To test the remaining activity of a component of the reaction mixture, saturating concentrations of the other two enzyme components were added, and the mixture was assayed for the ability to oxidize o-cresol to 3-methylcatechol. For example, the hydroxylase activity present in 10 ml of reaction mixture was assayed with additions of fresh small component (4 mM) and reductase (5 mM); to assay reductase activity, hydroxylase (1 mM) and small component (4 mM) were added; for the small component, hydroxylase (1 mM) and reductase (5 mM) were included. The activities of the enzyme components from the control reactions (without TCE) were also determined in this manner, and these activities were set at 100%. The percent activity remaining for each component from the inactivation reactions was then determined by comparison to the activity of the control. Analytical methods. Volatile compounds were detected with a Varian Aerograph Series 1400 gas chromatograph equipped with a flame ionization detector and a 0.1% AT-1000 Graphpac-GC 80/100 column (Alltech Associates [6-ft by 1/8-in. column]). The column temperature was 1808C, and the injector and detector temperatures were 2508C. Stoichiometries and retention times were determined by comparison of peak heights determined from standard curves with those of authentic compounds. Radiolabeled CO2 production was determined by venting the headspace of reaction vials into 2 N NaOH traps. Aliquots of the NaOH were added to scintillation fluid and analyzed for radioactivity. Nonradiolabeled carbon monoxide production was determined by carboxyhemoglobin formation (1). Acidic products were separated by HPLC on a Bio-Rad Aminex HPX-87H organic acid column (solvent, 10% acetonitrile in 0.01 N H2SO4; flow rate, 0.6 ml/min) and compared to authentic standards (12). The retention times obtained by monitoring the A210 were as follows: trichloroacetic acid, 6.5 min; dichloroacetic acid, 9.3 min; glyoxylate, 10.2 min; glycolate, 13.0 min; formate, 14.6 min; and chloral hydrate, 20.4 min. Fractions were collected at 20-s intervals and analyzed for radioactivity by scintillation counting. Products were quantified by measuring the radioactivity associated with each peak. Peaks of radioactivity were compared to the retention times of authentic standards. Incorporation of radiolabel into cell material or enzyme components was also determined. In whole-cell assays, cells were pelleted by centrifugation, washed once in 50 mM phosphate buffer (pH 7.0), and resuspended in fresh phosphate buffer. Aliquots of the washed cells were added to 7 ml of scintillation fluid, mixed, and counted for radioactivity. For reconstituted enzyme assays, aliquots of the reaction mixture were transferred to clean Eppendorf tubes, and the protein was precipitated with 5% trichloroacetic acid. The protein pellet was resuspended in 100 mM MOPS (pH 7.0) and analyzed for radioactivity. Chemicals and reagents. [U-14C]toluene (specific activity, 56.2 mCi/mmol) and [1,2-14C]TCE (specific activity, 23.3 mCi/mmol) were obtained from Sigma Radiochemicals. TCE, 3-methylcatechol, and o-cresol were obtained from Aldrich (Milwaukee, Wis.). Chloral hydrate, MOPS, glyoxylate reductase, formate dehy-

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FIG. 1. Water-soluble products of the oxidation of [14C]TCE by B. cepacia G4 analyzed by a chromatograph (HPLC) fitted with an organic acid column and scintillation counting. (A) Standard organic acid and aldehyde products. (B) Supernatant from 30-min incubation with [14C]TCE. (C) Supernatant from 30min incubation containing [14C]TCE and 10 mM glyoxylate.

drogenase, superoxide dismutase, and catalase were obtained from Sigma. All other chemicals used were reagent grade or better.

J. BACTERIOL.

In vitro TCE oxidation: requirements and kinetics. The three toluene 2-monooxygenase components, added singly and in combination, were analyzed for the ability to oxidize [14C] TCE to nonvolatile products. Two combinations of enzyme components, with the addition of NADH, were capable of oxidizing TCE. The complete enzyme system, or a combination of the hydroxylase and reductase, gave detectable levels of oxidation products (Table 1). The hydroxylase and the reductase alone oxidized TCE at only 10% the level of the fully reconstituted enzyme system, similar to the ratio previously observed with toluene as the substrate (28). The addition of reagents that scavenge reactive oxygen species did not diminish the formation of nonvolatile products, suggesting that the products were formed from an enzymatic oxidation of the substrate (Table 1). In fact, all of the additives tested, except mannitol, slightly increased the formation of nonvolatile products. The addition of superoxide dismutase and catalase, L-tryptophan, and the singlet oxygen scavenger L-histidine led to the greatest increase in the amount of nonvolatile products detected. HPLC analysis of the reaction mixtures revealed no additional organic acid products or chloral hydrate. The addition of catalase and superoxide dismutase to the reaction mixtures had a protective effect on the toluene 2-monooxygenase enzyme system incubated with NADH and O2 in the absence of organic substrate (data not shown). The formation of reactive oxygen species is often generated by oxygenases via uncoupling of NADH and substrate oxidation (2). Consistent with this was the observation that 2.5 mM NADH was utilized for each 1 mM TCE oxidized. The steady-state kinetic parameters for TCE oxidation were determined with purified toluene 2-monooxygenase components. The apparent Vmax and Km were 37 nmol/min mg hydroxylase component21 and 12 mM, respectively. This corresponds to a catalytic constant (kcat) of 8 min21 and a kcat/Km of 1 3 104 s21 M21 based on the hydroxylase a2b2g2 component. A measurement of the apparent kinetic constants was rendered difficult by the concomitant enzyme inactivation occurring over the first 5 min (described below). Assuming a firstorder loss of enzyme activity at the highest concentration of TCE assayed (2 mM), the Vmax, or kcat, is underestimated by at most 12%, with only a small effect on the Km. Products of in vitro TCE oxidation. HPLC analysis of the supernatant of 30-min enzyme-[14C]TCE incubation mixtures

TABLE 1. Effect of reaction mixture composition on nonvolatile product formation from TCEa Reaction mixture

RESULTS Initial whole-cell studies. TCE was incubated with intact cells of B. cepacia G4 expressing toluene 2-monooxygenase to gain insight into the potential enzyme reaction products. Nelson and coworkers reported that toluene-grown B. cepacia G4 transformed radiolabeled TCE to radiolabeled CO2, watersoluble product(s), and incorporated cell material (25). In the present study, the radiolabeled water-soluble product(s) were further analyzed by HPLC with an organic acids column. The single radioactive peak coeluted at 10.2 min with authentic glyoxylic acid (Fig. 1). In a 30-min incubation, radioactivity associated with the 10.2-min peak constituted 27% of the total radiolabel added to the reaction mixture. This decreased to 18% after 1 h. Addition of 10 mM nonradiolabeled glyoxylic acid increased recovery of radioactivity in the 10.2-min peak to 45% of the total [14C]TCE added.

Concn of nonvolatile products (nmol)

Complete (H, SC, R)b............................................................. 10.2 Complete 2 NADH................................................................ ,0.1 H, R........................................................................................... 0.96 SC, R......................................................................................... ,0.1 H alone ..................................................................................... ,0.1 NADH alone ............................................................................ ,0.1 TCE alone ................................................................................ ,0.1 Complete 1 40 mM mannitol ............................................... 10.0 Complete 1 2 mM L-tryptophan........................................... 11.4 Complete 1 10 mM L-histidine............................................. 12.1 Complete 1 SOD and catalasec ............................................ 13.2 a Reactions were performed as described in “Steady-state kinetic parameters of TCE oxidation” in Materials and Methods and were incubated for 15 min. b Complete denotes 1 mM hydroxylase, 2 mM small component, and 2 mM reductase, respectively. c SOD, superoxide dismutase. Concentrations of SOD and catalase were 100 U/ml.

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FIG. 2. Recovery of radioactivity from the oxidation of [14C]TCE by purified enzyme for 30 min with and without glyoxylate reductase or formate dehydrogenase. Reaction mixtures were analyzed on an HPLC organic acid column. (A) Reaction mixture containing toluene 2-monooxygenase only. (B) Reaction mixture containing toluene 2-monooxygenase plus glyoxylate reductase. (C) Reaction mixture containing toluene 2-monooxygenase plus formate dehydrogenase. Reaction conditions are as described in Materials and Methods.

on an organic acid column revealed two major radiolabeled products that comigrated with authentic glyoxylate and formate (Fig. 2A). The radiolabeled products eluting at 10.2 and 14.6 min were positively identified as glyoxylate and formate with the enzymes glyoxylate reductase and formate dehydrogenase, respectively. Glyoxylate reductase and NADH shifted the radiolabeled peak from 10.2 min to 13 min, the elution time of authentic glycolate (Fig. 2B). Addition of formate de-

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hydrogenase and NAD1 to a duplicate aliquot resulted in the disappearance of the radiolabeled peak at 14.6 min (Fig. 2C) and complete recovery of the radioactivity as 14CO2 (data not shown). Carbon monoxide was the only major volatile product (Table 2). No radiolabeled carbon dioxide was detected, indicating that the carbon monoxide formed was not further oxidized by the enzyme under the conditions used. The overall stoichiometry, determined after 90% oxidation of TCE, showed (i) 43% of the radiolabel was present as water-soluble products, (ii) 10% was associated with the protein fraction of the reaction mixture, and (iii) 41% was accounted for as CO in the volatile fraction (Table 2). The water-soluble products were accounted for as 10% of the oxidized TCE being glyoxylic acid and 21% being formic acid. Analysis of duplicate reactions at 7 and 15 min revealed the same 2.2:1.0 formate/glyoxylate ratio (data not shown). Determination of TCE epoxide formation. The products detected above suggested the formation of an epoxide intermediate that could be trapped in solution by reaction with 4-(pnitrobenzyl)pyridine (12, 16, 24). During the steady-state phase of a reaction time course, approximately 10% of the TCE oxidized in the enzymatic reaction could be trapped as TCE epoxide (data not shown). In a parallel experiment with purified soluble methane monooxygenase as a positive control, 13% of the TCE oxidized was trapped as TCE epoxide. This result is similar to the amount of TCE epoxide detected previously (12). In that study, 95% of the TCE oxidation products were reported to be derived from TCE epoxide. The relatively inefficient trapping of TCE epoxide in both enzymatic reactions, and as previously observed with cytochrome P-450 monooxygenase, is likely due to the short half-life (10 to 20 s) of TCE epoxide in solution under the conditions used in these experiments (24). Control experiments consisting of the reaction mixtures without addition of NADH or enzyme gave no detectable reactions with 4-(p-nitrobenzyl)pyridine. Covalent labeling of enzyme components. The protein-associated [14C]TCE products were examined further. Analysis of a reaction mixture containing all three enzyme components and [14C]TCE, but no NADH, revealed only trace amounts of radioactivity associated with the protein fraction. Complete reaction mixtures containing NADH were analyzed by SDSPAGE and quantitative determination of the radiolabel bound to each separated polypeptide. The results in Table 3 indicate that the hydroxylase and reductase components contained an equal density of label, but the small component contained a fourfold-higher density of radiolabel. Inactivation of the enzyme components. To investigate the kinetic course of TCE-dependent enzyme inactivation, the enzyme system was incubated with NADH, with and without TABLE 2. Product formation during the oxidation of [14C]TCEa Product

% of TCE oxidizedb

Carbon monoxide ...................................................................... 41.0 6 9.0 Formate ...................................................................................... 21.0 6 1.8 Glyoxylate................................................................................... 10.0 6 0.8 Dichloroacetic acid.................................................................... ,1.0 Trichloroacetic acid................................................................... ,1.0 Chloral ........................................................................................ ,1.0 Labeled protein ......................................................................... 12.0 6 1.8 a Products of the oxidation of radiolabeled TCE were determined as described in the text. Water-soluble products were analyzed with an HPLC organic acid column. Fractions were collected every 20 s and analyzed for radioactivity. b Determined for reactions in which 90% of the substrate was oxidized.

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TABLE 3. Radioactivity associated with the toluene 2-monooxygenase components after [14C]TCE oxidationa No. of cpm/As in trialb:

Component 1

2

Hydroxylase subunits a b g

0.022 0.022 0.030

0.015 0.015 0.017

Small component

0.096

0.050

Reductase

0.018

0.015

a

Radiolabeled enzyme components were analyzed by SDS-PAGE as described in Materials and Methods. b No. of CPM/As denotes radioactivity in counts per minute divided by the As of the protein component. The As for each enzyme component was calculated by the equation As 5 6.3 (Mw)0.73 (4).

TCE, and then assayed for the remaining enzyme activity at several time points. Figure 3 shows that significant time-dependent enzyme inactivation occurred. Seventy percent of the enzyme activity was lost after 30-min reactions with mixtures containing TCE and NADH, compared to a loss of only 25% in the absence of TCE. No loss of activity was detected in control reaction mixtures incubated with TCE alone (data not shown). L-cysteine protected the enzyme system during TCE turnover (Fig. 3). The loss of activity was similar to that of the control reaction mixture without TCE, suggesting that activity loss under the conditions used was due to the interception of a diffusible reactive intermediate(s). The degree of inactivation of each individual enzyme component was investigated by adding back saturating amounts of the other two enzyme components to determine the remaining activity of the third component. Aliquots of the TCE- and NADH-containing reaction mixtures from the experiment shown in Fig. 3 were assayed in this way. Figure 4 shows that the three enzyme components were inactivated to various extents, but the reductase component showed the greatest diminution in activity, retaining only 25 to 30% of that of the control. The cytochrome c reduction activity of the reductase was also decreased (data not shown). Addition of cysteine

FIG. 3. Time course of inactivation of the toluene 2-monooxygenase enzyme system during the oxidation of TCE. All reaction mixtures contain 3.5 mM NADH, superoxide dismutase, catalase, and 2 mM TCE, unless noted. 3, control (no TCE); Ç, complete mixture; E, complete mixture plus 10 mM cysteine. Reaction conditions are described in Materials and Methods.

FIG. 4. Inactivation of the components of the toluene 2-monooxygenase enzyme system. Aliquots of reaction mixtures from the experiment shown in Fig. 3 were tested at 30 min for activity with the substrate o-cresol. Supplementation of the reaction mixtures with saturating amounts of the reported components allowed for the determination of the remaining activity of the third component. The activities remaining in the control sample incubated without TCE were set at 100%. Solid bars, control (no TCE); hatched bars, complete mixture; open bars, complete mixture plus 10 mM cysteine.

resulted in a significant protection of the reductase; in comparison with the other two components, the activity remaining was then 75% that of the control. The hydroxylase component was not significantly protected by cysteine, and the small component was protected only slightly. DISCUSSION There is widespread interest in using B. cepacia G4 and derivative strains for bioremediation of TCE and related chlorinated solvents (10, 18, 19, 40), and this has created an imperative for research with purified enzyme components to identify the reaction products. For example, methane monooxygenase makes chloral as one of its oxidation products (12), and that compound is significantly more toxic than TCE (15, 34). Previously, only an in vivo study had been conducted with B. cepacia G4, and that showed substoichiometric production of carbon dioxide and unidentified water-soluble products (25). Toluene 2-monooxygenase was recently purified to homogeneity and found to consist of a flavo-iron-sulfur NADH-oxidoreductase, a small chromophoreless component, and a hydroxylase component that contains two binuclear-iron centers (28). In the presence of NADH and molecular oxygen, toluene 2-monooxygenase oxidizes toluene to ortho-cresol and orthocresol to 3-methylcatechol. Physiologically, it functions to activate the aromatic ring for a subsequent ring-fission reaction, allowing B. cepacia G4 to grow on toluene as its sole source of carbon and energy. Among known multicomponent microbial oxygenases, toluene 2-monooxygenase resembles soluble methane monooxygenase structurally. However, the substrate specificity of the two enzyme systems differ markedly. Toluene 2-monooxygenase does not oxidize methane (27). Conversely, methane monooxygenase oxidizes toluene to benzyl alcohol, ortho-cresol, meta-cresol, and para-cresol. The ratios of products differ depending on the source of enzyme (22, 39). Both enzymes are

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FIG. 5. Scheme for the oxidation of TCE by toluene 2-monooxygenase. T2M, toluene 2-monooxygenase. The spontaneous reactions of TCE epoxide (15) yielded the stable oxidation products shown on the far right.

implicated in TCE oxidation in vivo (11, 29), and methane monooxygenase has been studied with TCE in vitro (12). The latter enzyme oxidizes TCE predominantly to TCE epoxide, which spontaneously decomposes to dichloroacetic acid, glyoxylate, formate, and carbon monoxide in the enzyme reaction mixture. Synthetic TCE epoxide decomposes, in aqueous solution, to the same products (12, 16, 24). In this study, the major stable products of TCE oxidation by toluene 2-monooxygenase were shown to be carbon monoxide, formate, and glyoxylate (Fig. 5). Our results indicate that TCE epoxide is the precursor to the stable products. While the trapping reaction is substochiometric because of the instability of TCE epoxide (aqueous half-life of about 10 s), the percentages trapped from toluene 2-monooxygenase and methane monooxygenase reaction mixtures were similar, suggesting that TCE epoxide is the major or only direct oxidation product released by toluene 2-monooxygenase. Failure to detect dichloroacetate might be due to efficient trapping of its reactive precursor, dichloroacetyl chloride, by enzyme components. It is advantageous for bioremediation purposes that toluene 2-monooxygenase does not produce chloral, in contrast to all other monooxygenases investigated that produce it in variable proportions. In fact, chloral hydrate is the major product of the oxidation of TCE by mammalian cytochrome P-450 monooxygenases, although different isozymes appear to yield different fractional yields (24). Methane monooxygenase produces chloral at 6% of the total product yield (12). Chloral is proposed to derive from a reaction intermediate that can partition to either the epoxide or an aldehyde; the latter formation is accompanied by chloride migration yielding the trichloromethyl group (12, 24). It has been argued that enzymatic chloral formation does not derive from isomerization of TCE epoxide, and the failure to detect the former compound in this study is consistent with that idea. The various ratios of chloral produced by cytochrome P-450 monooxygenases are suggested to be due to subtle differences in active site environments that lead to differences in partitioning of the reaction intermediate between epoxide and aldehyde products. The lack of chloral formation by toluene 2-monooxygenase is explained by either the complete partitioning of a similar intermediate entirely to the epoxide or to the reaction proceeding through a different intermediate. The most unexpected finding, given previous in vivo experiments that suggested enzyme stability (11, 32), was that toluene 2-monooxygenase was covalently labeled and inactivated during the course of TCE oxidation in vitro. These data are consistent with the product data that suggested the generation of reactive intermediates such as TCE epoxide, formyl chloride, and glyoxyl chloride. The formation of covalent TCE adducts with all protein components was directly observed with [14C]TCE, and kinetic studies were also consistent with enzyme acylation. The addition of cysteine to reaction mixtures

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markedly protected the reductase component, but not the hydroxylase component, against inactivation. These data suggest inactivation by both diffusible intermediates and species that react before they come off the hydroxylase surface. Further protein chemistry studies are required to delineate the precise events leading to enzyme inactivation. The inactivation events discussed above are distinguishable from the slow NADH-mediated enzyme inactivation that occurred in the absence of TCE. This slow inactivation could be largely prevented through the addition of superoxide dismutase and catalase, suggesting the inactivation was mediated by diffusible reactive oxygen species. This type of inactivation has been observed with other oxygenases (2). This study is the first report of TCE-dependent inactivation of toluene 2-monooxygenase and differs from observations with whole cells that led to the conclusion that TCE-oxidizing activity was not lost during substrate turnover (11, 26). This is also in contrast to observations with soluble methane monooxygenase and toluene dioxygenase in which inactivation occurred both in vitro and in vivo (12, 20, 29, 37). The lack of inactivation in B. cepacia G4 during the in vivo oxidation of TCE suggests that the intracellular environment protects the toluene 2-monooxygenase enzyme system in some way. For example, this could be due to the presence of physiological thiols or other enzymes, such as epoxide hydrolase, that alter the fate of the primary oxidation product, TCE epoxide. Intracellular transformation of TCE epoxide, via enzymatic or nonenzymatic mechanisms, might also underlie the significantly higher level of glyoxylate observed in vivo compared to that seen in vitro. These possibilities are currently under investigation. ACKNOWLEDGMENTS We thank M. Shields, University of West Florida, for the gift of B. cepacia G4; Brad Wallar and John Lipscomb, University of Minnesota, for the gift of the purified soluble methane monooxygenase components; and Mike Logan, Rutgers University, for help in analyzing the kinetic data. This research was supported by cooperative agreement EPA/CR 820771-01-0 from the Environmental Protection Agency (to L.P.W.). REFERENCES 1. Ahr, H. J., L. J. King, W. Nastainczyk, and V. Ullrich. 1980. The mechanism of chloroform and carbon monooxide formation from carbon tetrachloride by microsomal cytochrome P-450. Biochem. Pharmacol. 29:2855–2861. 2. Bernhardt, F. H., and H. Kuthan. 1981. Dioxygen activation by putidamonooxin. The oxygen species formed and released under uncoupling conditions. Eur. J. Biochem. 120:547–555. 3. Brusseau, G. A., H.-C. Tsein, R. S. Hanson, and L. P. Wackett. 1990. Optimization of trichloroethylene oxidation by methanotrophs and the use of a colorimetric assay to detect soluble methane monooxygenase activity. Biodegradation 1:19–29. 4. Creighton, T. E. 1980. Proteins: structures and molecular properties, 2nd ed. W. H. Freeman and Company, New York. 5. Dabrock, B., M. Keßeler, B. Averhoff, and G. Gottschalk. 1994. Identification and characterization of a transmissible linear plasmid from Rhodococcus erythropolis BD2 that encodes isopropylbenzene and trichloroethylene catabolism. Appl. Environ. Microbiol. 60:853–860. 6. Ensign, S. A., M. R. Hyman, and D. J. Arp. 1992. Cometabolic degradation of chlorinated alkenes by alkene monooxygenase in a propylene-grown Xanthobacter strain. Appl. Environ. Microbiol. 58:3038–3046. 7. Ensley, B. D. 1991. Biochemical diversity of trichloroethylene metabolism. Annu. Rev. Microbiol. 45:283–299. 8. Epstein, J., R. W. Rosenthal, and R. J. Ess. 1955. Use of g-(4-nitrobenzyl) pyridine as analytical reagent for ethyleneimines and alkylating agents. Anal. Chem. 27:1435–1439. 9. Ewers, J., D. Freier-Schro¨der, and H.-J. Knackmuss. 1990. Selection of trichloroethylene (TCE) degrading bacteria that resist inactivation by TCE. Arch. Microbiol. 154:410–413. 10. Folsom, B. R., and P. J. Chapman. 1991. Performance characterization of a model bioreactor for the biodegradation of trichloroethylene by Pseudomonas cepacia G4. Appl. Environ. Microbiol. 57:1602–1608.

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11. Folsom, B. R., P. J. Chapman, and P. H. Pritchard. 1990. Phenol and trichloroethylene degradation by Pseudomonas cepacia G4: kinetics and interactions between substrates. Appl. Environ. Microbiol. 56:1279–1285. 12. 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. 13. Gibson, D. T., G. J. Zylstra, and S. Chauhan. 1990. Biotransformations catalyzed by toluene dioxygenase from Pseudomonas putida F1, p. 121–132. InS. Silver, A. M. Chakrabarty, B. Iglewski, and S. Kaplan (ed.), Pseudomonas: biotransformations, pathogenesis, and evolving biotechnology. American Society for Microbiology, Washington, D.C. 14. Goeptar, A. R., J. N. M. Commandeur, B. van Ommen, P. J. van Bladeren, and N. P. E. Vermeulen. 1995. Metabolism and kinetics of trichloroethylene in relation to toxicity and carcinogenicity. Relevance of the mercapturic acid pathway. Chem. Res. Toxicol. 8:3–21. 15. Goldenthal, E. I. 1971. A compilation of LD50 values in newborn and adult animals. Toxicol. Appl. Pharmacol. 18:185–207. 16. Henschler, D., W. R. Hoos, H. Fetz, E. Dallmeier, and M. Metzler. 1979. Reactions of trichloroethylene epoxide in aqueous systems. Biochem. Pharmacol. 28:543–548. 17. Hur, H.-G., M. J. Sadowsky, and L. P. Wackett. 1994. Metabolism of chlorofluorocarbons and polybrominated compounds by Pseudomonas putida G786(pHG-2) via an engineered metabolic pathway. Appl. Environ. Microbiol. 60:4148–4154. 18. Krumme, M. L., K. N. Timmis, and D. F. Dwyer. 1993. Degradation of trichloroethylene by Pseudomonas cepacia G4 and the constitutive mutant strain G4 5223 PR1 in aquifer microcosms. Appl. Environ. Microbiol. 59: 2746–2749. 19. Landa, A. S., E. M. Sipkema, J. Weijma, A. A. C. M. Beenackers, J. Dolfing, and D. B. Janssen. 1994. Cometabolic degradation of trichloroethylene by Pseudomonas cepacia G4 in a chemostat with toluene as the primary substrate. Appl. Environ. Microbiol. 60:3368–3374. 20. Li, S., and L. P. Wackett. 1992. Trichloroethylene oxidation by toluene dioxygenase. Biochem. Biophys. Res. Commun. 185:443–451. 21. Lipscomb, J. D. 1994. Biochemistry of the soluble methane monooxygenase. Annu. Rev. Microbiol. 48:371–399. 22. Liu, Y. 1995. Ph.D. thesis. University of Minnesota, Minneapolis. 23. Maltoni, C., and G. Lefimine. 1974. Carcinogenicity bioassays of vinylchloride. I. Research plan and early results. Environ. Res. 7:387–396. 24. Miller, R. E., and F. P. Guengerich. 1982. Oxidation of trichloroethylene by liver microsomal cytochrome P-450: evidence for chlorine migration in a transition state not involving trichloroethylene oxide. Biochemistry 21:1090– 1097. 25. Nelson, M. J. K., S. O. Montgomery, E. J. O’Neill, and P. H. Pritchard. 1986. Aerobic metabolism of trichloroethylene by a bacterial isolate. Appl. Environ. Microbiol. 52:383–384.

J. BACTERIOL. 26. Nelson, M. J. K., S. O. Montgomery, W. R. Mahaffey, and P. H. Pritchard. 1987. Biodegradation of trichloroethylene and involvement of an aromatic biodegradative pathway. Appl. Environ. Microbiol. 53:949–954. 27. Newman, L. M. 1995. Ph.D. thesis. University of Minnesota, St. Paul. 28. Newman, L. M., and L. P. Wackett. 1995. Purification and characterization of toluene 2-monooxygenase from Burkholderia cepacia G4. Biochemistry 34: 14066–14076. 29. 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. 30. Rannug, U., A. Johansson, C. Ramel, and C. A. Wachtmeister. 1974. The mutagenicity of vinyl chloride after metabolic activation. Ambio 3:194–197. 31. Segal, I. H. 1975. Enzyme kinetics. Behavior and analysis of rapid equilibrium and steady-state enzyme systems. John Wiley and Sons, New York. 32. Shields, M. S., and M. J. Reagin. 1992. Selection of a Pseudomonas cepacia strain constitutive for the degradation of trichloroethylene. Appl. Environ. Microbiol. 58:3977–3983. 33. Stanier, R. Y., N. J. Palleroni, and M. Doudoroff. 1966. The aerobic pseudomonads: a taxonomic study. J. Gen. Microbiol. 43:159–271. 34. Vagnarelli, P. A., A. Desario, and L. DeCarli. 1990. Aneuploidy induced by chloral hydrate detected in human keratinocytes with the Y97 probe. Mutagenesis 5:591–592. 35. Vogel, T. M., and P. L. McCarty. 1985. Biotransformation of tetrachloroethylene to trichloroethylene, dichloroethylene, vinyl chloride, and carbon dioxide under methanogenic conditions. Appl. Environ. Microbiol. 49:1080– 1083. 36. Wackett, L. P., and D. T. Gibson. 1988. Degradation of trichloroethylene by toluene dioxygenase in whole-cell studies with Pseudomonas putida F1. Appl. Environ. Microbiol. 54:1703–1708. 37. Wackett, L. P., and S. R. Householder. 1989. Toxicity of trichloroethylene to Pseudomonas putida F1 is mediated by toluene dioxygenase. Appl. Environ. Microbiol. 55:2723–2725. 38. Wackett, L. P., M. J. Sadowsky, L. M. Newman, H.-G. Hur, and S. Li. 1994. Metabolism of polyhalogenated compounds by a genetically engineered bacterium. Nature 368:627–629. 39. Wilkins, C., H. Dalton, C. J. Samuel, and J. Green. 1994. Further evidence for multiple pathways in soluble methane-monooxygenase-catalysed oxidations from the measurement of deuterium kinetic isotope effects. Eur. J. Biochem. 226:555–560. 40. Winkler, J., K. N. Timmis, and R. A. Snyder. 1995. Tracking the response of Burkholderia cepacia G4 5223-PR1 in aquifer microcosms. Appl. Environ. Microbiol. 61:448–455. 41. Young, R. B., M. Orcutt, and P. B. Blauwiekel. 1980. Quantitative measurement of protein mass and radioactivity in N9N-diallyltartardiamide crosslinked polyacrylamide slab gels. Anal. Biochem. 108:202–206.