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• W2015547178 abstract "2-Oxo-1,2-dihydroquinoline 8-monooxygenase, which catalyzes the NADH-dependent oxygenation of 2-oxo-1,2-dihydroquinoline to 8-hydroxy-2-oxo-1,2-dihydroquinoline, is the second enzyme in the quinoline degradation pathway of Pseudomonas putida 86. This enzyme system consists of two inducible protein components, which were purified, characterized, and identified as reductase and oxygenase. The yellow reductase is a monomeric iron-sulfur flavoprotein (Mr, 38,000), containing flavin adenine dinucleotide and plant-type ferredoxin [2Fe-2S]. It transferred electrons from NADH to the oxygenase or to some artificial electron acceptors. The red-brown oxygenase (Mr, 330,000) consists of six identical subunits (Mr, 55,000) and was identified as an iron-sulfur protein, possessing about six Rieske-type [2Fe-2S] clusters and additional iron. It was reduced by NADH plus catalytic amounts of reductase. For monooxygenase activity, reductase, oxygenase, NADH, molecular oxygen, and substrate were required. The activity was considerably enhanced by the addition of polyethylene glycol and Fe2+. 2-Oxo-1,2-dihydroquinoline 8-monooxygenase revealed a high substrate specificity toward 2-oxo-1,2-dihydroquinoline, since none of 25 other tested compounds was converted. Based on its physical, chemical, and catalytic properties, we presume 2-oxo-1,2-dihydroquinoline 8-monooxygenase to belong to the class IB multicomponent non-heme iron oxygenases. 2-Oxo-1,2-dihydroquinoline 8-monooxygenase, which catalyzes the NADH-dependent oxygenation of 2-oxo-1,2-dihydroquinoline to 8-hydroxy-2-oxo-1,2-dihydroquinoline, is the second enzyme in the quinoline degradation pathway of Pseudomonas putida 86. This enzyme system consists of two inducible protein components, which were purified, characterized, and identified as reductase and oxygenase. The yellow reductase is a monomeric iron-sulfur flavoprotein (Mr, 38,000), containing flavin adenine dinucleotide and plant-type ferredoxin [2Fe-2S]. It transferred electrons from NADH to the oxygenase or to some artificial electron acceptors. The red-brown oxygenase (Mr, 330,000) consists of six identical subunits (Mr, 55,000) and was identified as an iron-sulfur protein, possessing about six Rieske-type [2Fe-2S] clusters and additional iron. It was reduced by NADH plus catalytic amounts of reductase. For monooxygenase activity, reductase, oxygenase, NADH, molecular oxygen, and substrate were required. The activity was considerably enhanced by the addition of polyethylene glycol and Fe2+. 2-Oxo-1,2-dihydroquinoline 8-monooxygenase revealed a high substrate specificity toward 2-oxo-1,2-dihydroquinoline, since none of 25 other tested compounds was converted. Based on its physical, chemical, and catalytic properties, we presume 2-oxo-1,2-dihydroquinoline 8-monooxygenase to belong to the class IB multicomponent non-heme iron oxygenases. The soil bacterium Pseudomonas putida 86 utilizes quinoline as sole source of carbon, nitrogen, and energy. The occurrence of the metabolites 2-oxo-1,2-dihydroquinoline, 8-hydroxy-2-oxo-1,2-dihydroquinoline, 8-hydroxycoumarin, and 2,3-dihydroxyphenylpropionic acid revealed that the degradation of quinoline proceeds in this organism via the so called “coumarin pathway”(1Schwarz G. Bauder R. Speer M. Rommel T.O. Lingens F. Biol. Chem. Hoppe-Seyler. 1989; 370: 1183-1189Google Scholar, 2Shukla O.P. Appl. Environ. Microbiol. 1986; 51: 1332-1342Google Scholar). Quinoline 2-oxidoreductase from this organism, catalyzing the initial nucleophilic attack of quinoline at C-2, yielding 2-oxo-1,2-dihydroquinoline, is investigated thoroughly(3Bauder R. Tshisuaka B. Lingens F. Biol. Chem. Hoppe-Seyler. 1990; 371: 1137-1144Google Scholar, 4Tshisuaka B. Kappl R. Httermann J. Lingens F. Biochemistry. 1993; 32: 12928-12934Google Scholar). However, no enzymatic study is available concerning the further conversions of 2-oxo-1,2-dihydroquinoline in the “coumarin pathway.” In this paper, we present the purification and characterization of 2-oxo-1,2-dihydroquinoline 8-monooxygenase. This novel two-component enzyme system is compared with other multicomponent enzymes belonging to the family of non-heme iron oxygenases. DEAE-cellulose DE-52 was obtained from Whatman (Maidstone, Great Britain); phenyl-Sepharose CL-4B, Sephacryl S-300, Superose 12 HR 10/30, and Superdex 200 prep grade HiLoad 16/60 were from Pharmacia Biotech (Freiburg, Germany); DEAE-Fractogel EMD was obtained from Merck (Darmstadt, Germany); HPLC columns were from Bischoff (Leonberg, Germany); YM-10 and YM-30 ultrafiltration membranes were purchased from Amicon (Witten, Germany); and PEG1 1The abbreviations used are: PEGpolyethylene glycolDCPIP2,6-dichlorophenol indophenolgavaverage value of the g-tensor gav = 1/3 (g + g + g)HPLChigh pressure liquid chromatographyINT2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyl-2H-tetrazolium chloride. 6000 and carrier ampholytes (Servalyt 3-6) were obtained from Serva (Heidelberg, Germany). Quinoline and 2-oxo-1,2-dihydroquinoline were a gift from Ruetgerswerke AG (Castrop Rauxel, Germany). 8-Hydroxy-2-oxo-1,2-dihydroquinoline was obtained by biotransformation.2 2B. Tshisuaka, unpublished results. All other chemicals and biochemicals were of the highest purity commercially available. polyethylene glycol 2,6-dichlorophenol indophenol average value of the g-tensor gav = 1/3 (g + g + g) high pressure liquid chromatography 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyl-2H-tetrazolium chloride. P. putida 86 was described by Schwarz et al.(5Schwarz G. Senghas E. Erben A. Schfer B. Lingens F. Hke H. Syst. Appl. Microbiol. 1988; 10: 185-190Google Scholar). The strain was grown aerobically at 30°C on mineral salts medium containing (g/liter) 4.3 Na2HPO4× 2 H2O, 1.15 KH2PO4, 1.0 NaCl, 0.4 MgSO4× 7 H2O, 0.05 FeSO4× 7 H2O, and 0.01 Na2MoO4× 2 H2O. After sterilization, 4 mM quinoline was added. For studies of enzyme induction, the medium contained (instead of quinoline) 2.8 mM 2-oxo-1,2-dihydroquinoline or glucose (2 g/liter) plus (NH4)2SO4 (1 g/liter). After 15 h of growth, cells were washed twice with 50 mM Tris-HCl buffer (pH 7.5) and suspended (adjusting A578 to 2.2) in fresh mineral salts medium containing (i) 2.8 mM 2-oxo-1,2-dihydroquinoline or (ii) 2.8 mM 2-oxo-1,2-dihydroquinoline plus 50 mg/liter chloramphenicol. Degradation of 2-oxo-1,2-dihydroquinoline was monitored spectrophotometrically at 323 nm. Large-scale growth of bacteria was carried out in a 100-liter fermenter with quinoline as substrate. Bacterial growth and concentrations of quinoline and 2-oxo-1,2-dihydroquinoline were measured spectrophotometrically according to Bauder et al.(3Bauder R. Tshisuaka B. Lingens F. Biol. Chem. Hoppe-Seyler. 1990; 371: 1137-1144Google Scholar). After 9, 23, 32, and 48 h of fermentation, bacteria were fed with 4 mM quinoline. After total consumption of quinoline and 2-oxo-1,2-dihydroquinoline, cells were harvested by centrifugation, washed twice with 50 mM Tris-HCl buffer (pH 7.5), and stored at −20°C. All purification steps were carried out at 4°C, except for fast protein liquid chromatography (DEAE-Fractogel EMD and Superose 12), which was performed at room temperature. All pH values given refer to room temperature. In Step 1, frozen cells (40 g, wet weight) were thawed in 40 ml of standard buffer (50 mM Tris-HCl, pH 7.3) and disrupted 5 min in time intervals of 0.5 s in a sonifier (model 450, Branson Inc., Danbury, CT) with maximal power. The suspension was centrifuged at 48,000 × g for 40 min. In Step 2, the supernatant was diluted with the same volume of standard buffer and loaded for anion-exchange chromatography onto a DEAE-cellulose DE-52 column (2.5 × 39 cm) equilibrated in standard buffer. The column was washed with the same buffer, and bound proteins were desorbed with a linear gradient (800 ml) of KCl (0-0.6 M) in standard buffer at a flow rate of 0.6 ml/min. This purification step largely separated the reductase from the oxygenase. In Step 3, fractions containing the reductase were combined and adjusted to 0.8 M (NH4)2SO4. For hydrophobic interaction chromatography, the solution was applied to a column of phenyl-Sepharose (2.5 × 5 cm) equilibrated in 50 mM Tris-HCl buffer, pH 7.5, containing 0.8 M (NH4)2SO4 and 1 mM dithiothreitol. The column was rinsed with 200 ml of the same buffer, and proteins were eluted with a linear gradient (300 ml) of 0.8-0 M (NH4)2SO4, 50-10 mM Tris-HCl (pH 7.5), 0-10% (v/v) ethanol, containing 1 mM dithiothreitol. The flow rate was 3 ml/min. The combined fractions with reductase activity were rinsed with 50 mM Tris-HCl buffer, pH 7.8, containing 1 mM dithiothreitol, by ultrafiltration with a YM-10 membrane. In Step 4, for a second anion-exchange chromatography, the concentrated protein solution was applied to a DEAE-Fractogel EMD column (1 × 14 cm) equilibrated in 50 mM Tris-HCl buffer, pH 7.8, containing 1 mM dithiothreitol. The column was rinsed with 3 ml of the same buffer, and a linear gradient (35 ml) of 0-0.5 M (NH4)2SO4 was applied at a flow rate of 1 ml/min. Active fractions were pooled and concentrated to 0.5 ml by ultrafiltration with a YM-10 membrane. In Step 5, for gel filtration, the protein solution was loaded onto Superose 12 (1.5 × 28 cm), which was equilibrated in 100 mM Tris-HCl buffer, pH 7.8, containing 1 mM dithiothreitol. Proteins were separated at a flow rate of 0.5 ml/min. Fractions containing the reductase were pooled and stored at −80°C. Preparation of crude extract (Step 1′) and chromatography on DEAE-cellulose DE-52 (Step 2′) were performed as described above for the reductase, except that a crude extract from 20 g (wet weight) of cells was used. In Step 3′, fractions from DEAE-cellulose DE-52 containing the oxygenase were pooled and adjusted to 0.6 M (NH4)2SO4 and 3% (w/v) glycerol. The solution was loaded on a column of phenyl-Sepharose (2.5 × 5 cm), equilibrated in 50 mM Tris-HCl buffer, pH 7.5, containing 0.6 M (NH4)2SO4, 3% (w/v) glycerol, and 0.1 mM 2-oxo-1,2-dihydroquinoline. The column was washed with the same buffer, and the oxygenase was eluted by omitting (NH4)2SO4 from the buffer. To avoid severe losses of oxygenase activity, fast performance of all steps demanding high (NH4)2SO4 concentrations was substantial (flow rate, 10 ml/min). In Step 4′, after concentrating the eluate by ultrafiltration (YM-30 membrane), it was passed through a Sephacryl S-300 gel filtration column (2.5 × 92 cm; flow rate, 0.5 ml/min) using standard buffer containing 50 mM KCl and 0.1 mM 2-oxo-1,2-dihydroquinoline. In Step 5′, for adsorption chromatography on hydroxyapatite, which was prepared by the method of Atkinson et al.(6Atkinson A. Bradford P.A. Selmes I.P. J. Appl. Chem. Biotechnol. 1973; 23: 517-529Google Scholar), fractions with oxygenase activity were combined and adjusted to pH 7.2 with 1 M acetate buffer, pH 5.0. Glycerol was included to a final concentration of 3% (w/v), and 1 M potassium phosphate buffer, pH 7.2, was added to a final concentration of 10 mM. The solution was applied to a column of hydroxyapatite (2.5 × 5 cm), equilibrated in 10 mM potassium phosphate buffer, pH 7.2, containing 3% (w/v) glycerol and 0.1 mM 2-oxo-1,2-dihydroquinoline. The column was washed with the same buffer. Proteins were eluted with a linear gradient (300 ml) of 10-300 mM potassium phosphate buffer, pH 7.2 (3% (w/v) glycerol and 0.1 mM 2-oxo-1,2-dihydroquinoline), at a flow rate of 0.8 ml/min. For reproducible results, hydroxyapatite was used only once. Fractions containing the oxygenase were pooled, concentrated by ultrafiltration with a YM-30 membrane, and stored at −80°C. Since the oxygenase lost 50% specific activity when stored in standard buffer for 3 days at 4°C, different stabilizers were tested: 10% (w/v) glycerol, 10% (v/v) ethanol, 0.2% Triton X-100, 0.1 mM EDTA, 1-10 mM mercaptoethanol, 1 mM glutathione, 0.1-1 mM dithiothreitol, 0.1-1 mM (NH4)2Fe(SO4)2, and 0.1 mM 2-oxo-1,2-dihydroquinoline. Further attempts to preserve enzyme activity were made by using other buffers (pH 6-8.5) or purification procedures such as precipitation with (NH4)2SO4, PEG, or ethanol, chromatography on Superdex 200, chelating Sepharose, calcium tartrate or Mono-Q (anion exchange), and preparative native gel electrophoresis. Progress in enzyme purification was monitored by SDS-polyacrylamide gel electrophoresis with 10% separating and 4% stacking gels(7Schgger H. von Jagow G. Anal. Biochem. 1987; 166: 368-379Google Scholar). The 1-mm slab gels were stained in 0.2% (w/v) Coomassie Blue R-250 in water/methanol/acetic acid (40:50:10). Analytical isoelectric focusing was performed in rehydrated gels as recommended by Pharmacia(8Schickle H. Westermeier R. Pharmacia Bulletin. Freiburg, Germany1987Google Scholar). The native molecular weight of the reductase was determined by gel filtration through Superose 12 in 50 mM Tris-HCl buffer, pH 7.8, containing 150 mM NaCl. The native molecular weight of the oxygenase was estimated by gel filtration on Superdex 200 in 100 mM Tris-HCl buffer, pH 7.5. The molecular weights of peptides under denaturing conditions were determined using SDS-polyacrylamide gel electrophoresis(7Schgger H. von Jagow G. Anal. Biochem. 1987; 166: 368-379Google Scholar). Absorption spectra were measured in cells of 1-cm path length at 25°C with a Uvicon 930 spectrometer (Kontron Instruments, Neufahrn, Germany). The flavin cofactor was extracted from the reductase by two methods. The component in 10 mM potassium phosphate buffer (pH 7.0) was boiled for 4 min or treated with trichloroacetic acid as described by Siegel(9Siegel L.M. Methods Enzymol. 1978; 53: 419-429Google Scholar). After removal of protein by centrifugation and ultrafiltration, the yellow filtrate was examined by UV-visible spectroscopy (ϵFAD, 450 nm, 11,300 M-1 cm-1(10Whitby L.G. Biochem. J. 1953; 54: 437-442Google Scholar)) and by reversed phase HPLC on a Lichrospher RP18 column according to Nielsen et al.(11Nielsen P. Rauschenbach P. Bacher A. Methods Enzymol. 1986; 122: 209-220Google Scholar). 0.2 M formic acid, 0.1 M ammonia/methanol (3:1) was used as eluent. Acid-labile sulfur was determined by the formation of methylene blue as described by Beinert(12Beinert H. Anal. Biochem. 1983; 131: 373-378Google Scholar). Bovine milk xanthine oxidase was used as a standard. Analysis of metals was done with an x-ray fluorescence spectrometer (System 77 Finnigan Int. Inc., Sunnyvale, CA). Protein concentrations were determined according to Bradford (13Bradford M.M. Anal. Biochem. 1976; 72: 248-254Google Scholar) with bovine serum albumin as standard. The activity of 2-oxo-1,2-dihydroquinoline 8-monooxygenase was measured at 25°C spectrophotometrically at 365 nm as NADH consumption (standard assay, ϵ365, 3,400 M-1 cm-1(14Bergmeyer H.U. Z. Klin. Chem. Klin. Biochem. 1975; 13: 507-508Google Scholar)) or polarographically as oxygen uptake with a Clark-type oxygen electrode (YSI4004, Yellow Springs Instrument Co., Yellow Springs, OH). The reaction was optimized for temperature, pH, and buffer as well as for the concentrations of buffer, PEG, Fe2+, NADH, and 2-oxo-1,2-dihydroquinoline. The 21 mM Tris-HCl buffer (pH 7.5) contained 7% PEG, 0.05 mM (NH4)2Fe(SO4)2, 0.25 mM NADH, and suitable amounts of reductase and oxygenase components. The reaction was started by the addition of 0.2 mM 2-oxo-1,2-dihydroquinoline. All estimations were corrected for NADH or oxygen consumption recorded in the absence of substrate. 1 unit of 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity was defined as the amount of enzyme that consumed 1 μmol of NADH or 1 μmol of oxygen per min. The enzyme activity was not proportional to the crude extract concentration in the assay mixture. To achieve a proportional relationship, the activity of the reductase was measured in the presence of an excess amount of the oxygenase component and vice versa. These components to be added had been enriched by the purification steps 1 and 2. They were stored at −80°C until use. Substrate consumption and product formation were monitored by HPLC (Nucleosil RP18 column, eluent: 30% (v/v) methanol) with UV-visible spectroscopic detection at 200-400 nm. Substrate conversion was further confirmed by thin layer chromatography of ethyl acetate extracts of the acidified assay mixtures (silica gel plates; solvent system, toluene/dioxane/acetic acid (72:16:1.6)(1Schwarz G. Bauder R. Speer M. Rommel T.O. Lingens F. Biol. Chem. Hoppe-Seyler. 1989; 370: 1183-1189Google Scholar)). Authentic 8-hydroxy-2-oxo-1,2-dihydroquinoline served as product reference in both assays. The reaction mixtures contained 1.1 units of reductase, 12.5 milliunits of oxygenase, 0.05 mM (NH4)2Fe(SO4)2, 2 mM NADH, and 2 mM 2-oxo-1,2-dihydroquinoline in 50 mM Tris-HCl buffer (pH 7.5). Transformation of other putative substrates, causing oxygen consumption, was monitored likewise. Enzyme activity under anaerobic conditions (achieved by repeated degassing and flushing with nitrogen) was examined by thin layer chromatography as described above. NADH:acceptor reductase activity was assayed at 25°C spectrophotometrically as reduction of the artificial electron acceptors INT (ϵ503, 19,300 M-1 cm-1(15Babson A.L. Babson S.R. Clin. Chem. 1973; 19: 766-769Google Scholar)), DCPIP (ϵ600, 21,000 M-1 cm-1(16Armstrong J.M. Biochim. Biophys. Acta. 1964; 86: 194-197Google Scholar)), cytochrome c (ϵ550, 21,000 M-1 cm-1(17Massey V. Biochim. Biophys. Acta. 1959; 34: 255-256Google Scholar)), and potassium hexacyanoferrate III (ferricyanide) (ϵ420, 1,020 M-1 cm-1(18Schellenberg K.A. Hellerman L. J. Biol. Chem. 1958; 231: 547-556Google Scholar)). The reaction mixture contained 27 mM Tris-HCl buffer (pH 7.5), 0.05 mM electron acceptor, and 0.25 mM NADH. The reaction was started by the addition of 5-20 μl of protein solution. 1 unit of NADH:acceptor reductase activity was defined as the amount of enzyme that reduced 1 μmol of electron acceptor per min. The effects of several putative inhibitors on NADH:DCPIP reductase activity and on 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity (standard assay) were examined in the absence of Fe2+. Assay mixtures were preincubated 2 min with each compound (0.5 mM). The protein concentrations were 14 μg/ml in the NADH:DCPIP reductase assay and 760 μg/ml in the standard assay. Cells of P. putida 86, grown on glucose as sole source of carbon and energy, did not metabolize 2-oxo-1,2-dihydroquinoline in the presence of chloramphenicol, an inhibitor of protein synthesis. Without chloramphenicol, these cells degraded 2-oxo-1,2-dihydroquinoline after a lag phase of 2 h. Cells, grown on 2-oxo-1,2-dihydroquinoline, converted this substrate immediately when added anew without being affected by the addition of chloramphenicol. Thus, the degradation of 2-oxo-1,2-dihydroquinoline was dependent on protein biosynthesis induced by 2-oxo-1,2-dihydroquinoline. Extracts of cells, grown on glucose or succinate, showed neither reductase nor oxygenase activity of 2-oxo-1,2-dihydroquinoline 8-monooxygenase. If cells were grown on 2-oxo-1,2-dihydroquinoline, a specific reductase activity of 1.24 units/mg protein and a specific oxygenase activity of 0.35 units/mg protein was measured in the cell extracts. Therefore, we propose that both components of 2-oxo-1,2-dihydroquinoline 8-monooxygenase in P. putida 86 are inducible by the substrate 2-oxo-1,2-dihydroquinoline. The results of a typical purification procedure are summarized in Table 1. The reductase was purified 108-fold in a yield of 3%. The oxygenase activity was enriched only 2.4-fold. However, as judged by SDS-polyacrylamide gel electrophoresis, a high enrichment of the oxygenase protein in relation to other cell proteins was evident (Fig. 1). A rough estimate by comparing the gels of the oxygenase and reductase suggests that the increase in specific activity should have been around 100-fold. We assume that the oxygenase lost activity during the purification procedure despite stabilizing it with glycerol and substrate. Other tested stabilizers or purification procedures were even less effective in preserving enzyme activity.TABLE I Open table in a new tab The preparations of reductase and oxygenase were nearly homogeneous as shown by SDS-polyacrylamide gel electrophoresis (Fig. 1). The native molecular weight of purified reductase was determined to 39,000 by gel filtration. Its molecular weight determined under denaturing and reducing conditions by SDS-polyacrylamide gel electrophoresis was 37,000. Thus, we propose the reductase to be a monomer. Gel filtration of the oxygenase resulted in a native molecular weight of 330,000. Since SDS-polyacrylamide gel electrophoresis of the oxygenase revealed only one protein band corresponding to a molecular weight of 55,000, we suggest that the oxygenase is composed of six identical subunits. The isoelectric point of the purified reductase was 4.6. The oxygenase, which was nearly homogeneous as judged by SDS-polyacrylamide gel electrophoresis, separated in isoelectric focusing as a cluster of bands within the range of pH 4.9-5.4. As observed by other authors(19Frater R. J. Chromatogr. 1970; 50: 469-474Google Scholar, 20Cann J.R. Methods Enzymol. 1979; 61: 142-147Google Scholar), the formation of multiple bands might be an artifact due to complex formation of proteins with carrier ampholytes. In the oxidized state, the reductase was yellow in color. The UV-visible absorption spectrum showed maxima at 271 and 456 nm, a broad double peak at 337 and 381 nm, and shoulders at about 427 and 550 nm (Fig. 2). The reductase was reduced and bleached by the addition of NADH. After boiling and centrifuging a reductase solution, the supernatant had an absorption spectrum (Fig. 2, inset) typical for a flavin. Solutions of purified oxygenase, as isolated in the oxidized state, had a red-brown color. The UV-visible absorption spectrum showed maxima at 280, 328, and 460 nm and a shoulder at 545 nm (Fig. 3). Boiling of the oxygenase resulted in the loss of color. The oxygenase as isolated was bleached by chemical reduction with dithionite or by enzymatical reduction with NADH and catalytic amounts of reductase. The addition of NADH alone did not reduce the oxygenase. This finding indicated that the reductase, which was reduced directly by NADH, mediated the electron transfer from NADH to the terminal oxygenase component. Analysis by HPLC revealed an amount of 0.7 mol of FAD/mol of reductase. Spectrophotometrical determination of the FAD content resulted in a ratio of 0.9 mol of FAD/mol of reductase. Average contents of 1.7 ± 0.3 g atom iron and 1.6 ± 0.15 g atom acid-labile sulfur were determined per mol of reductase. 1 mol of oxygenase contained 13.1 ± 1.8 g atom iron and 9.1 ± 0.9 g atom acid-labile sulfur. The absorption spectra (Figure 2:, Figure 3:) indicate that the iron is not bound to a heme-like structure but is part of iron-sulfur clusters. Electron paramagnetic resonance studies revealed that a plant-type ferredoxin [2Fe-2S] cluster with gav = 1.95 is present in the reductase component, whereas the oxygenase component harbors Rieske-type [2Fe-2S] clusters (gav = 1.89).3 3B. Rosche, S. Fetzner, F. Lingens, W. Nitschke, and A. Riedel, submitted for publication. We propose that the reductase contains one FAD and one plant-type ferredoxin [2Fe-2S] cluster and that the oxygenase includes six Rieske-type [2Fe-2S] clusters (i.e. one [2Fe-2S] per subunit) and additional iron. Reductase, oxygenase, NADH, molecular oxygen, and substrate were required for 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity. Polarographic determinations of O2 uptake with limiting concentrations of either 2-oxo-1,2-dihydroquinoline or NADH revealed a 1:1:1 stoichiometry for 2-oxo-1,2-dihydroquinoline:O2:NADH. From the highest specific activities under conditions of the standard assay, turnover numbers of 1235 min-1 (reductase) and 320 min-1 (oxygenase) were calculated. No catalytic activity was detectable under anaerobic conditions. The pH optimum of the enzyme activity was pH 7.5. The optimal temperature was in the range of 25-30°C. The reductase component of 2-oxo-1,2-dihydroquinoline 8-monooxygenase showed NADH:acceptor reductase activity not only toward the oxygenase component of this enzyme system but also toward some artificial electron acceptors. Reductase with a specific activity of 40 units/mg, measured in the presence of the oxygenase component (standard assay), showed specific activities of 53, 89, 404, and 501 units/mg with INT, DCPIP, cytochrome c, and ferricyanide, respectively. The oxygenase showed no oxidoreductase activity with these acceptors. In the DCPIP reductase assay, the reductase would reach only 1.3% activity if NADH was replaced by NADPH. The addition of Fe2+ and FAD led to an increase in 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity (Table 2). We suggest that these additives might supplement cofactors lost during purification. FMN did not replace FAD, and NADPH was less effective as electron donor than NADH (Table 2). A preincubation of the enzyme components with Fe2+ and FAD (30 min, 4°C) did not further enhance activity.TABLE II Open table in a new tab None of the metal cations Mg2+, Ca2+, Fe3+, Co2+, and Ni2+ (50 μM each) substituted Fe2+ in its enzyme-activating effect. However, the addition of 50 μM Mn2+, Zn2+, or Cu2+ resulted in 14, 56, or 100% loss of activity, respectively, compared to the activity without any supplementary metal salt. In contrast to the 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity, the NADH:DCPIP reductase activity was not increased by the addition of Fe2+ or FAD. The NADH:DCPIP reductase was inhibited by Zn2+ and Cu2+ (33 and 100% inhibition), but it was not affected by Mn2+. 2-Oxo-1,2-dihydroquinoline (0.2 mM) did not influence the NADH:DCPIP reductase activity. A surprisingly strong activating effect on 2-oxo-1,2-dihydroquinoline 8-monooxygenase was observed by the addition of PEG to the enzyme assay. In the absence of PEG, the assay was nonlinear, initially showing only 14% of activity, which slowly increased to 38% within 12 min but never reached 100% as determined in the presence of PEG (Table 2). However, NADH:DCPIP reductase activity was diminished in the presence of PEG (67% residual activity). Substances that modify sulfhydryl groups (p-hydroxymercuribenzoate, N-ethylmaleimide, and iodoacetate, 0.5 mM each) affected both NADH:DCPIP reductase activity (98, 66, and 43% inhibition) and the activity of the complete monooxygenase system (97, 30, and 23% inhibition). Metal chelating agents (EDTA, 4,5-dihydroxy-1,3-benzene disulfonic acid, and 1,10-phenanthroline, 0.5 mM each) did not influence the NADH:DCPIP reductase activity, but the complete enzyme system was inhibited (12, 14, and 95%). 8-Hydroxyquinoline, 8-hydroxy-2-oxo-1,2-dihydroquinoline, and coumarin (0.1 mM each) caused substrate-dependent oxygen consumption without being converted. No oxygen consumption was observed with quinoline, 2-/8-monochloroquinoline, 2-/8-monomethylquinoline, quinoline 2-/8-monocarboxylic acid, 4-/5-/6-/7-monohydroxyquinoline, 2,4-/2,6-dihydroxyquinoline, isoquinoline, 1-oxo-1,2-dihydroisoquinoline, pyridine, acridine, indole, carbazole, hypoxanthine, 2-hydroxynaphthalene, and anthraquinone (0.1 mM each). An inducible two-component enzyme system, which catalyzes the NADH-dependent monooxygenation of 2-oxo-1,2-dihydroquinoline to 8-hydroxy-2-oxo-1,2-dihydroquinoline, was purified from P. putida 86. The enzyme system was termed 2-oxo-1,2-dihydroquinoline 8-monooxygenase with the systematic name 2-oxo-1,2-dihydroquinoline, NADH:oxygen oxidoreductase (8-hydroxylating). 2-Oxo-1,2-dihydroquinoline 8-monooxygenase revealed a high substrate specificity toward 2-oxo-1,2-dihydroquinoline, since none of 25 other compounds tested was converted. However, 8-hydroxyquinoline, 8-hydroxy-2-oxo-1,2-dihydroquinoline, and coumarin caused substrate-dependent oxygen consumption without being attacked. We assume that these compounds served as pseudosubstrates, uncoupling electron transfer from substrate hydroxylation with concomitant production of H2O2. This effect has been reported for some other oxygenases, e.g. orcinol hydroxylase(21Ribbons D.W. Ohta Y. FEBS Lett. 1970; 12: 105-108Google Scholar), salicylate hydroxylase(22White-Stevens R.H. Kamin H. J. Biol. Chem. 1972; 247: 2358-2370Google Scholar), or 4-methoxybenzoate monooxygenase(23Bernhardt F.-H. Erdin N. Staudinger H. Ullrich V. Eur. J. Biochem. 1973; 35: 126-134Google Scholar). Since exogenously added H2O2 decayed spontaneously under our test conditions, even in absence of any protein, H2O2 consequently was undetectable in the enzyme assay. A first hint that 2-oxo-1,2-dihydroquinoline 8-monooxygenase is a multicomponent enzyme system was the nonproportional relationship between enzyme activity and protein concentration. The specific enzyme activity decreased with protein dilution instead of remaining constant. Since contact of the two soluble enzyme components is a prerequisite for monooxygenase activity, a delayed binding may explain the nonlinear time dependence of activity in the absence of PEG (Table 2). The mechanism of enzyme activation by PEG might be based on strengthening the component contact due to its hygroscopic property. To profit from this activating effect, other multicomponent enzyme systems should be tested for the influence of PEG. Fig. 4 presents a model for the electron transport chain from NADH to the substrate hydroxylating terminal oxygenase. NADH transmits two electrons simultaneously as hydride, whereas iron-sulfur centers are restricted to one-electron reactions. The function of the reductase is to mediate this two-electron/one-electron transformation by its flavin cofactor. The oxygenase contains besides its Rieske-type iron-sulfur clusters additional iron. The necessity of iron for the catalytic function of the oxygenase component is demonstrated by the fact that metal chelating agents inhibited 2-oxo-1,2-dihydroquinoline 8-monooxygenase activity, while NADH:acceptor reductase activity was not affected. This agrees with the result that added ferrous iron did not increase NADH:acceptor reductase activity, whereas the complete enzyme reaction was accelerated. Therefore, we propose that ferrous iron is a weakly associated cofactor of the oxygenase component, which is abstracted easily by metal chelating agents or during enzyme purification and which may be replaced by exogenously added ferrous iron. According to our data, 2-oxo-1,2-dihydroquinoline 8-monooxygenase belongs to the group of non-heme iron multicomponent oxygenases, which contain both iron-sulfur clusters and additional iron in their oxygenase component. In contrast to flavin-containing single component monooxygenases or cytochrome P-450 multicomponent monooxygenases, which activate dioxygen by FAD or heme-bound iron, respectively(24Sariaslani F.S. Crit. Rev. Biotechnol. 1989; 9: 170-257Google Scholar), non-heme iron oxygenases are supposed to activate dioxygen by protein-bound iron(25Mason J.R. Cammack R. Annu. Rev. Microbiol. 1992; 46: 277-305Google Scholar). In the case of the oxygenase component of the O-demethylating 4-methoxybenzoate monooxygenase (putidamonooxin), mononuclear non-heme iron was demonstrated to mediate the electron transfer from a Rieske-type [2Fe-2S] cluster to molecular oxygen, thus activating the dioxygen as iron-peroxo complex for the electrophilic attack of the organic substrate(26Adrian W. Bernhardt F.-H. Bill E. Gersonde K. Heymann E. Trautwein A. Twilfer H. Hoppe-Seyler's Z. Physiol. Chem. 1980; 361: 211Google Scholar, 27Bill E. Bernhardt F.-H. Trautwein A.X. Eur. J. Biochem. 1981; 121: 39-46Google Scholar). This model might be representative for all known multicomponent non-heme iron oxygenases, which contain Rieske-type iron-sulfur clusters and mononuclear iron in their terminal oxygenase component. In Table 3(28Locher H.H. Leisinger T. Cook A.M. Biochem. J. 1991; 274: 833-842Google Scholar, 29Locher H.H. Leisinger T. Cook A.M. J. Bacteriol. 1991; 173: 3741-3748Google Scholar, 30Bernhardt F.-H. Pachowsky H. Staudinger H. Eur. J. Biochem. 1975; 57: 241-256Google Scholar, 31Bernhardt F.-H. Heymann E. Traylor P.S. Eur. J. Biochem. 1978; 92: 209-223Google Scholar, 32Markus A. Krekel D. Lingens F. J. Biol. Chem. 1986; 261: 12883-12888Google Scholar, 33Schweizer D. Markus A. Seez M. Ruf H.H. Lingens F. J. Biol. Chem. 1987; 262: 9340-9346Google Scholar, 34Batie C.J. LaHaie E. Ballou D.P. J. Biol. Chem. 1987; 262: 1510-1518Google Scholar, 35Brunel F. Davison J. J. Bacteriol. 1988; 170: 4924-4930Google Scholar, 36Fetzner S. Mller R. Lingens F. J. Bacteriol. 1992; 174: 279-290Google Scholar, 37Haak B. Fetzner S. Lingens F. J. Bacteriol. 1995; 177: 667-675Google Scholar, 38Yamaguchi M. Fujisawa H. J. Biol. Chem. 1978; 253: 8848-8853Google Scholar, 39Yamaguchi M. Fujisawa H. J. Biol. Chem. 1980; 255: 5058-5063Google Scholar, 40Yamaguchi M. Fujisawa H. J. Biol. Chem. 1982; 257: 12497-12502Google Scholar, 41Bnz P.V. Cook A.M. J. Bacteriol. 1993; 175: 6467-6475Google Scholar, 42Sauber K. Frhner C. Rosenberg G. Eberspcher J. Lingens F. Eur. J. Biochem. 1977; 74: 89-97Google Scholar, 43Geary P.J. Saboowalla F. Patil D. Cammack R. Biochem. J. 1984; 217: 667-673Google Scholar, 44Axcell B.C. Geary P.J. Biochem. J. 1975; 146: 173-183Google Scholar, 45Subramanian V. Liu T.-N. Yeh W.-K. Gibson D.T. Biochem. Biophys. Res. Commun. 1979; 91: 1131-1139Google Scholar, 46Subramanian V. Liu T.-N. Yeh W.-K. Narro M. Gibson D.T. J. Biol. Chem. 1981; 256: 2723-2730Google Scholar, 47Subramanian V. Liu T.-N. Yeh W.-K. Serdar C.M. Wackett L.P. Gibson D.T. J. Biol. Chem. 1985; 260: 2355-2363Google Scholar, 48Erickson B.D. Mondello F.J. J. Bacteriol. 1992; 174: 2903-2912Google Scholar, 49Ensley B.D. Gibson D.T. J. Bacteriol. 1983; 155: 505-511Google Scholar, 50Haigler B.E. Gibson D.T. J. Bacteriol. 1990; 172: 457-464Google Scholar, 51Haigler B.E. Gibson D.T. J. Bacteriol. 1990; 172: 465-468Google Scholar, 52Takizawa N. Kaida N. Torigoe S. Moritani T. Sawada T. Satoh S. Kiyohara H. J. Bacteriol. 1994; 176: 2444-2449Google Scholar), 2-oxo-1,2-dihydroquinoline 8-monooxygenase is compared with some of these multicomponent non-heme iron oxygenases. Batie et al.(53Batie C.J. Ballou D.P. Correll C.C. Mller F. Chemistry and Biochemistry of Flavoenzymes. CRC Press, Boca Raton, FL1991: 543-556Google Scholar) grouped them into the three classes I, II, and III, based on the number of protein components and on the redox centers involved in electron transport from NAD(P)H to the terminal oxygenase component. At least two redox centers, a flavin and a [2Fe-2S] center, are involved in this electron transfer. A second [2Fe-2S] center is possible. The redox centers can be located on one or two protein components, thus constituting two- or three-component enzyme systems. Class I enzymes are two-component enzyme systems with a flavin (FMN in class IA, FAD in class IB) and a plant-type ferredoxin [2Fe-2S] in the reductase. In class II enzymes, the flavin (always FAD) and the [2Fe-2S] center are located on separate components. The [2Fe-2S] center is a plant-type ferredoxin (class IIA) or a Rieske-type center (class IIB). Class III enzymes contain FAD and plant-type ferredoxin in the reductase as well as a Rieske-type center in a second component. According to this classification, 2-oxo-1,2-dihydroquinoline 8-monooxygenase belongs to class IB enzyme systems, as do 2-halobenzoate 1,2-dioxygenase(36Fetzner S. Mller R. Lingens F. J. Bacteriol. 1992; 174: 279-290Google Scholar, 37Haak B. Fetzner S. Lingens F. J. Bacteriol. 1995; 177: 667-675Google Scholar), benzoate 1,2-dioxygenase(38Yamaguchi M. Fujisawa H. J. Biol. Chem. 1978; 253: 8848-8853Google Scholar, 39Yamaguchi M. Fujisawa H. J. Biol. Chem. 1980; 255: 5058-5063Google Scholar, 40Yamaguchi M. Fujisawa H. J. Biol. Chem. 1982; 257: 12497-12502Google Scholar), and toluate 1,2-dioxygenase(54Harayama S. Rekik M. Timmis K.N. Mol. & Gen. Genet. 1986; 202: 226-234Google Scholar). However, the oxygenase components of these three class IB enzymes are heteromultimers, whereas 2-oxo-1,2-dihydroquinoline 8-monooxygenase is a homomultimer like the class IA enzymes listed in Table 3.TABLE III Open table in a new tab 2-Oxo-1,2-dihydroquinoline 8-monooxygenase also bears structural resemblance to other multicomponent non-heme iron monooxygenases like toluene 4-monooxygenase(55Whited G.M. Gibson D.T. J. Bacteriol. 1991; 173: 3010-3016Google Scholar), phenol hydroxylase(56Powlowski J. Shingler V. J. Bacteriol. 1990; 172: 6834-6840Google Scholar), or xylene monooxygenase(57Suzuki M. Hayakawa T. Shaw J.P. Rekik M. Harayama S. J. Bacteriol. 1991; 173: 1690-1695Google Scholar, 58Shaw J.P. Harayama S. Eur. J. Biochem. 1992; 209: 51-61Google Scholar), but the oxygenase components of these enzyme systems apparently do not possess iron-sulfur centers. Whether 2-oxo-1,2-dihydroquinoline 8-monooxygenase is a true monooxygenase or whether the monohydroxylated product is due to spontaneous dehydration of an unstable 7,8- or 8,8a-dihydrodiol of 2-oxo-1,2-dihydroquinoline, thus implying a dioxygenase reaction, is uncertain. Consequently, it remains to be determined whether 8-hydroxy-2-oxo-1,2-dihydroquinoline is an in vivo intermediate in the “coumarin pathway” of quinoline degradation by P. putida 86. However, no intermediate was detected during the enzyme reaction in vitro, even not under moderate conditions, taking into account the instability of a putative dihydrodiol intermediate. A 7,8-dihydrodiol was formed from the substrate analog 2-chloroquinoline by resting cells of P. putida 86(59Fetzner S. Vogler B. Lingens F. FEMS Microbiol. Lett. 1993; 112: 151-158Google Scholar). This activity is now shown to be independent from 2-oxo-1,2-dihydroquinoline 8-monooxygenase because the latter enzyme system did not convert 2-chloroquinoline. This paper demonstrates once more the diversity of hydroxylating enzymes encountered in quinoline-degrading bacteria. Schwarz et al.(1Schwarz G. Bauder R. Speer M. Rommel T.O. Lingens F. Biol. Chem. Hoppe-Seyler. 1989; 370: 1183-1189Google Scholar) described two pathways of quinoline degradation. In the first step of both pathways, quinoline is converted to 2-oxo-1,2-dihydroquinoline, catalyzed by quinoline 2-oxidoreductase, which incorporates a hydroxyl group deriving from water(3Bauder R. Tshisuaka B. Lingens F. Biol. Chem. Hoppe-Seyler. 1990; 371: 1137-1144Google Scholar, 60Peschke B. Lingens F. Biol. Chem. Hoppe-Seyler. 1991; 372: 1081-1088Google Scholar). Whereas the quinoline 2-oxidoreductases even from distantly related bacteria exhibit far-reaching similarities(3Bauder R. Tshisuaka B. Lingens F. Biol. Chem. Hoppe-Seyler. 1990; 371: 1137-1144Google Scholar, 60Peschke B. Lingens F. Biol. Chem. Hoppe-Seyler. 1991; 372: 1081-1088Google Scholar, 61Hettrich D. Peschke B. Tshisuaka B. Lingens F. Biol. Chem. Hoppe-Seyler. 1991; 372: 513-517Google Scholar), 4 4S. Schach, B. Tshisuaka, S. Fetzner, and F. Lingens, submitted for publication. the oxygenases catalyzing the next degradation step attack the same substrate (2-oxo-1,2-dihydroquinoline) at different positions. The 5,6-dioxygenating enzyme of Comamonas testosteroni4 consequently totally differs from the enzyme system of P. putida 86, hydroxylating at C-8 (this paper). The capabilities of enzymes, performing nucleophilic as well as electrophilic regio- (and stereo-)selective hydroxylations should give a stimulus to their industrial use for biotransformations(62Holland H.L. Organic Synthesis with Oxidative Enzymes. VCH Publishers, Inc., New York, NY1992Google Scholar). We are grateful to Dr. A. Riedel (Universitt Regensburg, Germany) for performing electron paramagnetic resonance spectroscopy. We thank Prof. Dr. Schreiber for recording and interpreting x-ray fluorescence spectra and K. Kapassakalis for technical assistance during fermentation." @default.
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• W2015547178 title "2-Oxo-1,2-dihydroquinoline 8-Monooxygenase, a Two-component Enzyme System from Pseudomonas putida 86" @default.
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