Eur. J. Biochem. 232, 536-544 (1995) 0 FEBS 1995

Quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase from Comamonas testosteroni 63 The first two enzymes in quinoline and 3-methylquinoline degradation

Susanne SCHACH, Barbara TSHISUAKA, Susanne FETZNER and Franz LINGENS

Institut fur Mikrobiologie der Universitat Hohenheim, Stuttgart, Germany

(Received 8 May 1995) - EJB 95 072414

The enzymes catalysing the first two steps of quinoline and 3-methylquinoline degradation by Comu- monus testosteroni 63 were investigated. Quinoline 2-oxidoreductase, which catalyses the hydroxylation of (3-methy1L)quinoline to (3-methyl-)2-oxo-l,2-dihydroquinoline, was purified to apparent homogeneity. The native enzyme, with a molecular mass of 360 kDa, is composed of three non-identical subunits (87, 32, and 22 kDa), occurring in a ratio of 1.16: 1 :0.83. Containing FAD, molybdenum, iron, and acid-labile sulfur in the stoichiometric ratio of 2: 2: 8 : 8, the enzyme belongs to the molybdo-ironhlfur flavoproteins. Molybdopterin cytosine dinucleotide is the organic part of the pterin molybdenum cofactor. Comparison of N-terminal amino acid sequences revealed similarities to a number of procaryotic molybdenum-con- taining hydroxylases. Especially the N-termini of the b-subunits of the quinoline 2-oxidoreductases from Comamonas testosteroni 63, Pseudomonas putidu 86, and Rhodococcus spec. B1, and of quinoline-4- carboxylic acid 2-oxidoreductase from Agrobacterium spec. 1 B showed striking similarities.

Further degradation of (3-methyl-)2-oxo-l ,Zdihydroquinoline proceeds via dioxygenation at the ben- zene ring, i.e. at 5,6-position [Schach, S., Schwarz, G., Fetzner, S. & Lingens, F. (1993) Bid. Chem. Hnppe-Seyler 374, 175- 1811. 2-0x0-I ,2-dihydroquinoline 5,6-dioxygenase was partially purified; NADH and oxygen are required for the reaction, and the enzymic activity is enhanced 1.5-fold by addition of Fez+ ions. Unexpectedly, this aromatic ring dioxygenase did not separate into distinct protein conipo- nents, but is apparently a single-component enzyme. The molecular mass was estimated to be about 260 kDa. 2-0~0-1,2-dihydroquinoline 5,6-dioxygenase is very thermolabile. However, dithioerythritol and low concentrations of substrate had a moderately stabilizing effect. 2-0x0-I ,2-dihydroquinoline 5,6- dioxygenase is inhibited by sulfhydryl-blocking agents, by metal-chelating agents, and by the flavin analogues quinacrine and acriflavin.

Keywords: quinoline 2-oxidoreductase; 2-0x0-I ,2-dihydroquinoline 5,6-dioxygenase; 5,6-dihydro-5,64- hydroxy-2-0x0-I ,2-dihydroquinoline ; aromatic ring dioxygenase; molybdenum-containing hydroxylase.

Several catabolic pathways of microbial degradation of quin- oline and quinoline derivatives are known (Schwarz et al., 1989; Tibbles et al., 1989; Bott et al., 1990; Hund et al., 1990; Schmidt et al., 1991; Schach et al., 1993; Riiger et al., 1993; Dembek et al., 1989; Grant and Al-Najjar, 1976; Shukla, 1986,

Correspondence to F. Lingens, Institut fur Mikrobiologie (250), Uni- versitat Hohenheim, Garbenstr. 30, D-70593 Stuttgart, Germany

Fax: +49 I1 1 459 2238. Abbreviations. (SH),Ery-ol, dithioerythritol ; INT, 2-(4-iodophenyl)-

3-(4-nitrophenyl)-5-phenyI-2ff-tetrazolium chloride. En7yrne.v. Carbon monoxide dehydrogenase, carbon monoxide :(ac-

ceptor) oxidoreductase (EC 1.2.99.2); isoquinoline I-oxidoreductase, isoquinoline:(acceptor)l-oxidoreductase (hydroxylating); nicotinic acid dehydrogenase, pyridine 3-carboxylic acid:(acceptor)6-oxidoreductase (hydroxylating) (EC 1.5.1.13); 2-0x0-I ,2-dihydroquInoline 5,6-dioxy- genase. 2-0x0- 1,2-dihydroquinoIine, NADH: oxygen 5,6-oxidoreductase (5,6-hydroxylating) (EC 1.14.12.-); quinaldic acid 4-oxidoreductase, quinoliiie-2-carboxyIic acid: (acceptor)4-oxidoreductase (hydroxylating) (EC 1 .-.-.-); quinaldine 4-oxidoreductase, 2-methylquinoline: (accep- tor)4-oxidoreductase (hydroxylating) (EC 1 ; quinoline 2-oxidore- ductase, quinoline:(acceptor)2-oxidoreductase (hydroxylating) (EC 1 ; quinoline-4-carboxylic acid 2-0xidoreductase, quinoline-4-car- boxylic acid : (acceptor)2-oxidoreductase (hydroxylating) (EC 1

Note. The N-terminal amino acid sequence data of the subunits of quinoline 2-oxidoreductase from Cornurnonas testosteroni have been deposited in thc PIR and Swiss-Prot protein sequence databases and are availabk under accession numbers P80464 ( n chain), P80465 (/,I chain), and P80466 (7 chain).

1989). Considering the degradation of quinoline, two divergent pathways were described, which both involve the formation of 2-0x0-I ,2-dihydroquinoline. Some pseudomonads further de- grade 2-0x0-I ,2-dihydroquinoline via the coumarin pathway by a monooxygenation at C8 (Rosche et al., 1995), formation of 8- hydroxycoumarin, and 2,3-dihydroxyphenylpropionate (Shukla, 1986, 1989; Schwarz et al., 1989). In Rhodococcu~ spec. B1 (Schwarz et al., 1989) and a Mnraxellu sp. (Grant and Al-Najjar, 1976), however, the degradation of 2-0x0-I ,2-dihydroquinoline has been suggested to proceed via 6-hydroxy- and the hypotheti- cal 5,6-dihydroxy-2-oxo-1,2-dihydroquinoline, which then un- dergoes meta-cleavage.

Our studies on the catabolism of 3-methylquinoline by Co- mamonas testosteroni 63 established 3-methyl-2-0x0-I ,2-dihy- droquinoline, 5,6-dihydroxy-3-methyl-2-oxo-1,2-dihydroquino- line, and 2,5,6-trihydroxy-3-methylpyridine as intermediates in the degradation pathway (Schach et a]., 1993). Additionally, 6-hydroxy-3-methyl-2-oxo-l.2-dihydroquinoline was isolated from the culture supernatant. However, the mechanism of 5,6- dihydroxy-3-methyl-2-oxo-1,2-dihydroquinoline formation re- mained obscure: either this compound is formed by two subse- quent monooxygenation steps with 6-hydroxy-3-methyl-2-0x0- 1,2-dihydroquinoline as intermediate, or a dioxygenase catalyses the formation of a cis-dihydrodiol, which upon dehydrogenation would yield 5,6-dihydroxy-3-methyI-2-oxo-1,2-dihydroquino- line. A spontaneous dehydration of the dihydrodiol would then

Schach et al. ( E m J . Biochem. 232) 537

account for the occurrence of 6-hydroxy-3-methyl-2-oxo-l,2-di- hydroquinoline. It was found that quinoline is degraded by C. testosteroni 63 via the same pathway as 3-methylquinoline.

Enzymic studies were performed to elucidate the mode of (3-methyl-)2-oxo-l,2-dihydroquinoline conversion. The novel aromatic ring dioxygenase involved is shown to be a 2-oxo-1,2- dihydroquinoline 5,6-dioxygenase, and some of its properties are discussed. Quinoline 2-oxidoreductase, catalysing the first step of (3-methyl-)quinoline degradation in C. testosteroni 63, was purified and characterized. This enzyme is compared with other oxidoreductases involved in the degradation of quinoline and derivatives that belong to the family of molybdo-irordsulfur fla- voproteins.

MATERIALS AND METHODS

Materials. Phenyl-Sepharose CL4B, Superdex 200 prep grade HiLoad 16/60, SuperoseTM 12 HR 10/30, and low-molecu- lar-mass standard for SDS/PAGE were from Pharmacia Biotech. DEAE-Fractogel EMD (1 cmX 14 cm) was from Merck. Nucleo- sil RP 18 ( 5 pm, 2.50 mmX4.6 mm) and LiChrospher RP 18 (5 pm, 250 mmX4.6 mm) were from Bischoff (Leonberg, Ger- many). Polyvinylidene difluoride transfer membrane was purchased from Millipore. Precoated silica plates SilG-UV,,, for TLC were from Macherey & Nagel. Quinoline 2-oxidoreductase from Pseudomonas putida 86 was purified as described by Tshi- suaka et al. (1993). 3-Methylquinoline was obtained from Lan- caster Synthesis (Eastgate, White Lund, Morecambe, England). Quinoline and 2-0~0-1,2-dihydroquinoline were a gift from Rutgerswerke AG (Castrop-Rauxel, Germany). 8-Hydroxy-2- oxo-1,2-dihydroquinoline was obtained by biotransforniation using P. putida 86 (Tshisuaka, unpublished results). 3-Methyl-2- 0x0- 1,2-dihydroquinoline and 6-hydroxy-2-0x0- 1,2-dihydroqui- noline were a gift from G. Schwarz. All other chemicals were of the highest purity commercially available.

Organism and culture conditions. The bacterial strain Comamonas testosteroni 63 and its growth conditions were de- scribed previously (Schach et al., 1993).

For large-scale growth, the bacteria were cultured in a 100-1 fermentor. Substrate consumption and metabolite formation were followed by TLC analysis (Schach et al., 1993). After 24, 38, 48, and 60 h of fermentation, the bacteria were fed with an additional portion of either quinoline or 3-methylquinoline; 2 h after the last substrate addition, the cells were separated by cen- trifugation and stored at -80°C until use.

Protein determination. Protein was determined according to Bradford (1976), with bovine serum albumin as standard.

Enzyme assays. Quinoline 2-oxidoreductase activity was determined photometrically by following the increase in absor- bance at 503 nm due to the substrate-dependent reduction of the monotetrazolium salt p-iodonitrotetrazolium violet (INT) to its red formazan = 19.3 mM-' cm-'; Babson and Babson, 1973). The standard assay mixture consisted of 750 pl INT ( 5 mM in distilled H20), 250 pl test buffer (0.2 M Tris/HC1 pH 8.0, 1 % Triton X-loo), 10 p1 quinoline or 3-methylquinoline (each 170 mM in ethanol), and 10 p1 enzyme solution. The reac- tion was started with substrate and was performed at 25°C. One unit of enzyme activity was defined as the amount catalyzing the formation of 1 pmol formazadmin.

Substrate specificity was determined with the standard assay modified as follows: SO p1 substrate to be tested (170 mM in ethanol) were added to 250 pl INT, 750 pI test buffer, and 10 p1 enzyme solution.

The activity of 2-0~0-1,2-dihydroquinoline 5,6-dioxygenase was assayed by measunng substrate-dependent oxygen con-

sumption with a Clark-type oxygen electrode (YSI4004, Yellow Springs Instrument Co., Yellow Springs OH). The standard as- say was optimized for the type of buffer, pH, and concentrations of ingredients. It contained in 20 mM Tris/HCl pH 7.3 : enzyme solution, 0.3 mM NADH, 0.2 mM (NH4),Fe(S0J2, and 0.15 mM substrate. One unit of enzyme was defined as the amount of enzyme that catalyzed the consumption of 1 pmol oxygedmin at 25°C.

The pH optimum of 2-oxo-l,2-dihydroquinoline 5,6-dioxy- genase was determined with the standard assay, replacing the test buffer by 20 mM sodium succinate in the pH range 5.0- 6.5, by 20 mM potassium phosphate in the pH range 6.0-7.2, and by 20 mM Tris/HCl in the pH range 7.3-8.5.

To test the optimal molarity of the buffer, the buffer used in the standard assay was replaced by 20, 50, and 100 niM potas- sium phosphate pH 6.5, 7.0, and 7.5 and by 20, 50, and 100 mM Tris/HCl pH 7.3, 7.5, 8.0, and 8.5.

In order to investigate the influence of metal cations on the activity of 2-0xo-l,2-dihydroquinoline 5,6-dioxygenase, (NH,),Fe(SO,), of the standard assay was replaced by other metal salts at a final concentration of 0.01 or 0.1 mM. The influ- ence of various salts (NaCl, KCl, (NH,),SO,, and CaCI,) was measured by adding them to the standard assay in a final con- centration of 0.2 M.

To test putative inhibitors for their concentration-dependent effect on the 2-0~0-1,2-dihydroquinoline 5,6-dioxygenase, the inhibitor was added to the standard assay mixture. After an incu- bation time of 2 min, the reaction was started by addition of substrate and the residual activity was determined. For testing the influence of putative inhibitors during prolonged incubation, the enzyme, dissolved in 20 mM Tris/HCl pH 7.3, was incubated together with the inhibitor at 4°C. Aliquots were taken after 2 and 4 h for performing the standard enzyme assay.

Substrate specificity of 2-0xo-l,2-dihydroquinoline 5,6-di- oxygenase was determined in the standard enzyme assay, replac- ing 2-0x0-I ,2-dihydroquinoline by various putative substrates ( 5 mM in isopropanol or S O mM Tris/HCl pH 7.5). The com- pounds tested were: quinoline, 1 -oxo-l,2-dihydroisoquinoline, naphthalene-2-01, 4-, 6- or 8-monohydroxy-2-oxo-l,2-dihy- droquinoline, acridine, coumarin, indole, carbazole, quinazoline, pyridine, hypoxanthine, 4-, 5-, 6- or 8-monohydroxyquinoline, quinoline-2-carboxylic acid, kynurenic acid, 2-methylquinoline, and 2- or 4-monochloroquinoline.

To test putative stabilizing agents for their effect on the 2- oxo-l,2-dihydroquinoline 5,6-dioxygenase, they were added to the protein solution. The enzyme activity was measured in the standard assay immediately and after incubation overnight at 4°C.

In order to investigate the thermostability, aliquots of en- zyme solution were incubated for 5 min at temperatures in the range of 30-65°C. The residual activity was measured in the standard assay.

Purification of quinoline 2-oxidoreductase and of 2-0x0- 1,2-dihydroquinoline 5,6-dioxygenase. All steps were per- formed at 4°C or in an ice bath, except for FPLC which was performed at room temperature.

Frozen cells (0.5 g wet mass/ml buffer) were suspended in 50 mM Tris/HCl pH 7.5 and sonified for 7 min with maximal power in time intervals of 0.5 s (Branson sonifier 450). The ho- mogenate was centrifuged for 60 min at 48000Xg. Proteins of the crude extract which were precipitated with (NH,),SO, (20% saturation) were discarded after centrifugation, and the superna- tant was heated to 45 "C for 5 min. After centrifugation, the solu- tion was loaded onto a phenyl-Sepharose column equilibrated in 50 mM Tris/HCl containing 20% (mass/vol.) (NH,),SO,. The column was washed with this buffer, then the buffer's content

538 Schach et al. (Eur: J. Biochem. 232)

Table 1. Purification of quinoline 2-oxidoreductase and of 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase from C. testosteroni 63. Starting mate- rial was 50 g cells.

Purification step Quinoline 2-oxidoreductase 2-0x0-I ,2-dihydroquinoline 5,6-dioxygenase - ___-

total total specific yield puri- total total specific yield puri- protein activity activity fication protein activity activity fication

1% U U/mg % -fold mg U U/mg % -fold

Extract 5200 2470 0.48 1 00 1 5200 111 0.02 100 1 Ammonium sulfate 4990 2115 0.44 88 0.9 4990 102 0.02 87 1 Heat treatmcnt 2150 1540 0.56 62 1.2 2150 71 0.03 66 1.5 Pheny I-Sepharose 320 1280 4 51 8.3 320 16 0.05 14 2.5 DEAE-Fractogel 3.3 15 4.5 0.6 9.4 56 I 0.125 0.7 6 Superdex 200 2.9 14 4.8 0.6 10 - - - - -

of (NH,),SO, was decreased to 10% (mass/vol.) before a linear gradient of 10% (NH,),SO, in SO mM Tris/HCI pH 7.5, to 0% (NH&SO, 10 mM Tris/HCl pH 7.3 (500 ml) was applied. Quin- oline 2-oxidoreductase and 2-0~0-1,2-dihydroquinoline 5,6-di- oxygenase coeluted in a final wash with 10 mM Tris/HCl pH 7.3. Active fractions were pooled, concentrated by ultrafiltra- tion with an Aniicon YM-100 membrane and applied to a DEAE-Fractogel column, equilibrated in 20 mM Tris/HCI pH 7.5, connected to a FPLC system. The column was washed with the equilibration buffer and both enzymes desorbed with a linear 75-ml gradient of 20-600 mM Tris/HCl pH 7.5. Fractions showing activities were pooled and concentrated by ultrafiltra- tion. The concentrate of the dioxygenase was stored at -80°C after glycerol (lo%, by vol.) had been added. For gel filtration, the concentrated quinoline 2-oxidoreductase was applied in 0.5-ml portions to a Superdex 200 column, equilibrated in 50 mM Tris/HCI pH 7.5. Proteins were separated at a flow rate of 60 ml/h. Active fractions were pooled, concentrated, and stored at -80°C.

Gel electrophoresis. The purity of quinoline 2-oxidoreduc- tase preparations was monitored by discontinuous non-denatur- ing polyacrylamide gels at high pH according to Hames (1990), containing 10% polyacrylamide in the resolving gel. Activity staining of quinoline 2-oxidoreductase was performed by incu- bating the gels in the standard assay mixture. Protein was stained with Coomassie brilliant blue R-250.

In order to determine the subunit composition and the sub- unit masses of quinoline 2-oxidoreductase, SDSPAGE was per- formed using 7.5%, lo%, and 12.5% separating gels (Schagger and von Jagow, 1987).

Molecular mass estimations. The molecular masses of the native enzymes were estimated by gel filtration on Superdex 200 and on Superose 12. Both columns were equilibrated in 50 mM Tris/HCl pH 7.3, containing 5 mM dithioerythritol [(SH),Ery-011 and 0.1 mM 2-0xo-l,2-dihydroquinoline as stabilizing additives. Proteins used for calibration were as follows: ferritin (440 kDa), glutamate dehydrogeilase (320 kDa), xanthine oxidase (283 kDa), catalase (230 kDa), and y-globulin (169 kDa).

Identification of the reaction product of 2-0xo-1,2-dihy- droquinoline 5,6-dioxygenase. In order to identify the product of the enzymic conversion of (3-methyl-)2-oxo-l,Z-dihydroqui- noline, 1 ml enzyme solution was incubated with 1 ml 3 mM substrate, 100 p1 10 mM NADH, and 100 pl 10 mM Fez+. Ali- quots were taken at appropriate intervals, and the reaction was stopped by boiling for 4 min. After removing the protein by cen- trifugation and ultrafiltration, the solution was examined by HPLC and TLC.

For TLC analysis, the samples were separated with toluene/ dioxane (312, by vol.) as solvent. For the detection and differen-

tiation of phenols and vicinal diols, TLC plates were sprayed with 2 % dichloroquinone 4-chloroimide in methanol, and metu- periodate/benzidine reagent (Cifonelli and Smith, 1954), respec- tively.

Analytical and preparative HPLC were performed on a Nucleosil RP 18 column with water/methanol (70/30. by vol.) as mobile phase (flow rate 0.7 ml/min). The spectral focus detector (Thermo Separation Products, California) connected to the HPLC system registered the ultraviolethisible spectra of the separated compounds in the range 200-400 nm. The samples were subjected to HPLC analysis (a) directly after ultrafiltration and (b) after acidification with HC1. 2-0~0-1,2-dihydroquinoline and 6-hydroxy-2-oxo-1,2-dihydroquinoline served as authentic references.

The optical activity of the reaction product was examined with a Perkin-Elmer polarimeter 241.

Metal, cofactor, and acid-labile sulfur determinations. All analyses were performed using two different concentrations of quinoline 2-oxidoreductase. The content of iron and molybde- num of quinoline 2-oxidoreductase was determined by X-ray fluorescence spectroscopy (system 77, Finnigan Internat. Inc., USA).

The flavin cofactor was extracted from the quinoline 2-oxi- doreductase by (a) treatment of the enzyme with trichloroacetic acid (final concentration lo%), and (b) boiling the enzyme for 5 min. Protein was removed by centrifugation and ultrafiltration, and the cofactor solution was analysed according to Nielsen et al. (1986) by HPLC on a LiChrospher RP-18 column at a flow rate of 0.8 ml/min with 0.1 M NH,, 0.2 M HCOOH in 25% (by vol.) methanol as eluent. The retention times of authentic FAD, FMN, and riboflavin were compared with the retention time ob- tained with the cofactor solution at 254 nm.

The identity of the nucleotide moiety in the pterin molybde- num cofactor was assessed as developed by Frunzke and Meyer (1991) and described by Hettrich et al. (1991) with AMP, CMP, GMP, and UMP as references.

Acid-labile sulfur was determined according to Beinert (1983), using milk xanthine oxidase as reference protein.

Absorption spectra. Protein absorption spectra were mea- sured with a Perkin-Elmer spectrophotometer UV/Vis lambda 12.

Determination of the amino-terminal protein sequences. The three subunits of the purified quinoline 2-oxidoreductase from C. testosteroni 63 were separated on a 10% SDSi polyacrylamide gel. The protein bands were transferred to a polyvinylidene difluoride transfer membrane by semi-dry electroblotting according to Kyhse-Andersen (1 984). Amino-ter- mind sequence analysis was accomplished by automated Edman

Schach et al. (Eur: J. Biochem. 232) 539

- 1 z u 0.8 F

0.6

0.4 D

s g n o

.; 0.2 4-

I c

0 10 20 30 40 50 60 70 g Fraction no. 2

Fig. 1. Chromatographic behaviour of quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase during anion-ex- change chromatography on DEAE-Fractogel. (-) Protein mea- sured by absorbance at 280 nm; (- - - - -) Tris/HCl gradient; (-) activ- ity of (A) quinoline 2-oxidoreductase and of (B) 2-oxo-l,2-dihydroqui- noline 5,6-dioxygenase.

1 2 3

Fig. 2. SDSFAGE of purified quinoline 2-oxidoreductases from R putidu 86 and C. testosteroni 63. Lane 1 , standard proteins (94 kDa, 67 kDa, 43 kDa, 30 kDa, 20.1 kDa, 14.4 kDa); lane 2, R putidu enzyme; lane 3, C. testosteroni enzyme.

microsequencing (Applied Biosystems protein sequencer model 476A).

RESULTS Purification of quinoline 2-oxidoreductase and of 2-0x0-1,2- dihydroquinoline 5,6-dioxygenase. Quinoline 2-oxidoreductase was purified from Comamonas testosteroni 63 in a five-step pro- cedure. Because of its instability, only partial purification of 2- 0x0-I ,2-dihydroquinoline 5,6-dioxygenase was achieved. The data of typical purification schemes are summarized for both enzymes in Table 1. The addition of (NH,),SO,, on the one hand, stabilized 2-0x0-I ,2-dihydroquinoline 5,6-dioxygenase in the heat treatment step, on the other hand, it was necessary for hydrophobic interaction chromatography. Fig. 1 illustrates the separation of quinoline 2-oxidoreductase and 2-0x0-I ,2-dihy- droquinoline 5,6-dioxygenase by means of anion-exchange chro- matography.

Physical, chemical, and catalytic properties of quinoline 2- oxidoreductase. Molecular mass and subunit composition. The molecular mass of the native quinoline 2-oxidoreductase, esti- mated by gel filtration, was 360 kDa. Polyacrylamide gel elec- trophoresis under reducing and denaturing conditions showed

2 .o

3 1.0 c a n L n

0.0 300

0.0 300 400 500 600

Wavelength [nml

Fig. 3. Absorption spectra of quinoline 2-oxidoreductase from C. tes- tosteroni 63. Curve 1, oxidized form (as isolated); curve 2, reduced with quinoline; curve 3, reduced with sodium dithionite.

Table 2. Substrate specificity of quinoline 2-oxidoreductases from C. testosteroni 63 and from E! putida 86.

Substrate Activity of enzyme from

C. testosteroni 63 R putidu 86

%

Quinoline 3-Methylquinoline 4-Methylquinoline 8-Methylquinoline 6-Hydroxyquinoline 4-Chloroquinoline 8-Chloroquinoline Isoquinoline

100 61 12 54 58

5 55 75

100 22 21 32 25

8 29 0

three protein bands corresponding to molecular masses of 87 kDa (a), 32 kDa (p), and 22 kDa ( y ) (Fig. 2). Based on these subunit masses, integration of a densitometric scan of a Coomas- sie-blue-stained gel revealed a molar subunit ratio of a/p/y = 1 .I 6 : 1 : 0.83.

Metal and cofactor analysis. Purified quinoline 2-oxidore- ductase contained I .6 mol molybdenum and 7.7 mol iron/mol enzyme. For acid-labile sulfur, an average value of 7.5 mol/mol enzyme was determined. The flavin cofactor was determined as FAD, amounting to 1.8 mol/mol enzyme. CMP, which was iden- tified in the protein free cofactor solution, is part of the pterin molybdenum cofactor. These data indicate a molybdo-iron/sulfur flavoprotein with molybdopterin cytosine dinucleotide as part of the pterin molybdenum cofactor.

Absorption spectra. The ultravioletlvisible spectra of homo- geneous quinoline 2-oxidoreductase (Fig. 3) were typical for a molybdo-ironhlfur flavoprotein. The oxidized enzyme (as iso- lated) is characterized by absorption maxima at 273 nm and 320 nm, an increase in absorbance in the range of 420-450 nm, and a shoulder at 550 nm. The enzyme was reduced by addition of substrate. The substrate binding spectrum is characterized by a decrease in absorbance in the region 360-580 nm and an increase above 600 nm. Reduction with sodium dithionite still further bleached the enzyme. The A28cJA450 ratio of the enzyme was 5.75, indicating its high purity (Coughlan, 1980). The ratio

540 Schach et al. (Eur: J. Riochem. 232)

a-Subunit

I ) A K S D V A E L K P R

2 ) M M K H E V V A L K q K S I G T

4) V E S I S A P K M V G Q A l P

6 ) A V V S Q D V A P P D G Q A E L N D

I \

P-Subunit

I ) M K F P A F A Y 7 R

2 M K F P A F S Y R A P A S L Q E V I Q ’ L A

3) M K A P A F q Y A 7 P A T L D E T F A

4) M R P F Q F I T P H S A A E A R E H L A

5) M ’ J N F A F L E P A T V A E A S Q M L A

y-Subunit

I ) M I Q A E K N P ? L

2) M Q A H E E S Q L M R I S A T 1 N G K P ’ J V F

3) M N E S ? E A ? A A ? ? V T I E V N

4) M R E V T I S V N G R P V T A E I D E R

5 ) M N Q V S I E L T V N G E E H Q V

6 ) M K F N L N G Q A V E F N G E P D T

7) M I E F I L N G Q P V ? V T E V P E ? A

Fig. 4. Comparison of amino-terminal protein sequences of subunits of several oxidoreductases. ( 1 ) Quinoline 2-oxidoreductase from C. tes- rosteroni 63 (this work) ; (2) quinoline 2-oxidoreductase from P. putith 86 and Khorlticoccu.s spec. B1 (Peschke and Lingens, 1991); (3) quino- line-4-carboxylic acid 2-oxidoreductase from Agrohacrerium spec. 1 B (unpublished) ; (4) quinaldine 4-oxidoreductase from Arthrohacter spec. Rii6la (de Beyer and Lingens, 1993); ( 5 ) quinaldic acid 4-oxidoreduc- tase from Pseudoinonus sp. AK-2 (unpublished); (6) quinaldic acid 4- oxidoreductase from Serrufia niurcescens 2CC-1 (Fetzner and Lingens, 1993) ; (7) isoquinoline 1 -oxidoreductme from Pseudumonus diminura 7 (Lehmarin et al., 1995).

A,,,JA,,,, of 3.0 is typical for an iron-sulfur/flavin ratio of 4 : 1 (Rajagopalan and Handler, 1964).

Amino-terminal protein sequences. The NH,-terminal amino acid sequences of the three different subunits of quinoline 2- oxidoreductase from C. testosteroni 63 were compared to the corresponding stretches of amino acids of other molybdenum- containing hydroxylases involved in the bacterial catabolism of quinoline (derivatives). The alignment revealed significant simi- larities, especially concerning the N-termini of the &subunits (Fig. 4).

Substrate specifcity. The substrate specificities of quinoline 2-oxidoreductase from C. testosteroni 63 and of quinoline 2- oxidoreductase from Pseudomonas putida 86 towards some quinoline derivatives are compared in Table 2. Quinoline 2-oxi- doreductase from C. testosteroni 63 shows a significantly higher activity with 3-methylquinoline than the quinoline 2-oxidore- ductase from f? putidu 86. C. testosteroni 63 had been isolated from activated sludge due to its ability to use 3-methylquinoline as sole source of carbon, nitrogen, and energy (Schach et al., 1993). The substrate usually used for growth was %methyl- quinoline. Nevertheless, it is more justified to name this new enzyme quinoline 2-oxidoreductase instead of 3-methyl- quinoline 2-oxidoreductase, because quinoline was converted with maximum activity.

Properties of 2-oxo-1,2-dihydroquinoline 5,idioxygenase. Product of the enzymic conversion. Aliquots of a large-scale re-

Wavelength [nml

Fig. 5. Absorption spectrum of 5,6-dihydro-5,6-dihydroxy-3-methyl- t-oxo-1,2-dihydroquinoline.

action mixture were separated by TLC. When sprayed with 2 % dichloroquinone 4-chloroimide, one nwabolite of the 2-oxo-l,2- dihydroquinoline conversion yielded a brown spot. The very same metabolite showed a yellow spot on a dark blue back- ground when sprayed with meta-periodatelbenzidine reagent. Both findings indicate a vicinal diol structure. When acidified samples of the reaction mixture after TLC were sprayed with these reagents, no positive reactions were observed.

The HPLC chromatogram of the neutral samples showed among other peaks one well resolved peak at 19 min with the ultraviolet spectrum depicted in Fig. 5. This substance was opti- cally active. HPLC analysis of the acidified samples revealed that the peak eluting at 19 min disappeared, and a new one at 12.7 min was observed. As judged by its absorption spectrum (Schwarz et al., 1989) and by comparison with the authentic reference, the latter compound was identified as 6-hydroxy-2- 0x0- 1,2-dihydroquinoline.

All these data suggest the formation of a dihydrodiol deriva- tive of 2-0x0-I ,2-dihydroquinoline. Since the degradation path- way proceeds via 5,6-dihydroxy-2-oxo-1,2-dihydroquinolirre (Schach et al., 1993), the 5,6-dihydrodiol (rather than the 6,7- dihydrodiol) of 2-0~0-1,2-dihydroquinoline seems plausible.

Physical properties. Gel filtration of the native enzyme on Superose 12 and Superdex 200 revealed molecular masses of 240 kDa and 280 kDa, respectively. During the purification pro- cedure, no separation of 2-oxo-l,2-dihydroquinoline 5,6-dioxy- genase into distinct protein components occurred, either by hy- drophobic interaction, anion-exchange chromatography, or gel filtration. Other methods tested for purification of the enzyme, for example adsorption chromatography with hydroxyapatite or with calcium tartrate, and chelating Sepharose charged with CU’+ or Zn”, did not indicate a multicomponent system either. In contrast to other aromatic ring dioxygenases which under such conditions completely separate into their components (e.g. Ensley et al., 1982; Fetzner et al., 1989; Romanov and Hau- singer, 1994), 2-0x0-l ,2-dihydroquinoline 5,6-dioxygenase is apparently a single-component enzyme.

Cutulytic properties. 2-0~0-1,2-dihydroquinoline 5,6-dioxy- genase activity was present i n cells grown on quinoline or on 3- methylquinoline only if they were harvested when the consump- tion of (3-methyl-)2-0~0-1,2-dihydroquinoline was in progress. Enzymic conversion of (3-methyl-)2-oxo-l,2-dihydroquinoline did not occur under anaerobic conditions. In vitro, the enzyme required NADH as electron donor, which could not be replaced by NADPH.

The activity of the enzyme was enhanced 1.5-fold by Fe2‘ ions. When Fez was added to the standard assay together with Fe”, the activity of 2-oxo-l,2-dihydroquinoline 5,6-dioxygen- ase did not reach the maximum activity measured with Fez+ alone. Cu’+ ions inhibited the enzyme completely. All other metal cations tested (CoZ+, Zn2+, Ca” , Ni*+, Mg2+, MnZ+) had no significant influence on enzyme activity.

Schach et al. (Eur: J. Biochem. 232) 541

Table 3. Concentration-dependent effects of various inhibitors on 2- oxo-1,2-dihydroquinoline 5,6-dioxygenase activity.

Reagent Concentration Residual activity

mM %

1,lO-Phenanthroline 0.01 73 0.1 37 0.5 18

Iodoacetate 1 1.5 2

61 44 0

4-Hydroxymercuribenzoate 0.01 71 0.02 68 0.1 0

Acriflavin 0.1 0.2 0.5

88 73 31

Quinacrine 0.1 100 0.2 92 0.5 85

The 2-0~0-1,2-dihydroquinoline 5,6-dioxygenase had a pH optimum of 7.3. The optimal molarity of all buffers tested was 20 mM.

The presence of salts conventionally used in ion-exchange or hydrophobic-interaction chromatography was disadvanta- geous for the enzyme activity. Addition of 0.2 M NaCl resulted in a 50% loss of activity. In the presence of 0.2 M KC1 and 0.2 M CaCI,, only 15% and 10% residual activity remained, respectively. However, 0.2 M (NH,),SO, caused a decrease in activity of only 12%, so that 88% of the initial activity re- mained.

Substrate specificity was tested with the purest preparation available. Besides 2-0~0-1,2-dihydroquinoline and 3-methyl-2- oxo-l,2-dihydroquinoline, only 8-hydroxyquinoline, 6-hydroxy- 2-0x0-1 ,2-dihydroquinoline, and 8-hydroxy-2-0x0-l ,2-dihy- droquinoline were converted. All other compounds tested did not elicit any oxygen consumption.

The dependence of enzyme activity on protein concentration was examined. In the range tested, the specific activity was inde- pendent of protein concentration. In contrast, in multicomponent enzyme systems consisting of dissociable protein components, the specific activity decreases with protein dilution. This typical feature is evident even in crude extracts (e.g. Ribbons, 1977; Ensley et al., 1982; Markus et al., 1984; Thurnheer et al., 1990).

Inhibitors. The inhibitory effects of metal chelating agents, thiol reagents, and of flavin analogues are summarized in Table 3. The effect of chelating agents during prolonged incuba- tion with enzyme is shown in Table 4.

All three types of reagents inhibited the enzyme. The weak chelators (sodium diethyldithiocarbaminate and EDTA) were in- hibitory only after a prolonged incubation time. In the short term, they even activated the 2-0~0-1,2-dihydroquinoline 5,6- dioxygenase.

Stability. 2-0~0-1,2-dihydroquinoline 5,6-dioxygenase was extremely thermolabile. Incubation at 30°C for 5 min caused a decrease in activity of 30%. When incubated for 5 min at 65"C, all activity was lost.

In order to stabilize 2-0~0-1,2-dihydroquinoline 5,6-dioxy- genase, several additives were tested. (SH),Ery-01, dithiothreitol and 2-oxo-l,2-dihydroquinoline (final concentration 0.1 mM) were found to stabilize the enzyme (Table 5) . No effect was found with 2% (mass/vol.) sucrose, 5 % (mass/vol.) glucose, or

Table 4. Effects of chelating agents on 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase activity after a prolonged incubation time.

Reagent Concen- Residual activity after tration

Oh 2 h 4 h

mM %

2,2'-Dipyridyl 1 83 70 64

Tiron 1 61 50 21

Sodium diethyldithio- 0.1 1 04 1 00 86 carbaminate 1 107 60 21

EDTA 0.1 90 85 50 1 116 90 29

Table 5. Effects of stabilizing additives on 2-oxo-1,2-dihydroquino- line 5,6-dioxygenase.

Additive Concen- Residual activity after tration

0 days 1 day

mM 7%

None 100 39

2-0~0-1,2-dihydroquinoline 1 95 50 0.1 102 70

Dithiothreitol 1 126 59 5 132 81

(SH),EV-ol 1 122 61 5 122 95

with 5 , 10 or 15% (by vol.) glycerol. Ethanol in a final concen- tration of 5 % (by vol.) decreased the enzyme activity to 44 %.

DISCUSSION

In Cornamonas testosteroni 63, (3-methyl-)quinoline degra- dation is initiated by hydroxylation at C2 (Fig. 6). Hydroxylation adjacent to the N heteroatom, an electron-deficient site, is a common mode of primarily attacking N heterocycles (e.g. Schwarz et al., 1989; Shukla, 1989; Ruger et al., 1993). Quino- line 2-oxidoreductase from C. testosteroni 63 resembles in many of its properties other molybdo-iron/sulfur flavoproteins catalys- ing a hydroxylation adjacent to the heteroatom of quinoline and its derivates (Bauder et al., 1990; Peschke and Lingens, 1991 ; Bauer and Lingens, 1992). Other enzymes of an analogous struc- ture are quinaldine 4-oxidoreductase from Arthrobacter spec. Rii6la (de Beyer and Lingens, 1993), quinaldic acid 4-oxidore- ductase from Pseudomonas sp. AK-2 (Sauter et al., 1993), car- bon monoxide oxidases of carboxydotrophic bacteria (Meyer et al., 1986; Johnson et al., 1990) and nicotinate dehydrogenase from Bacillus niacini (Nagel and Andreesen, 1990). Their com- mon features are the molecular masses of the native enzymes, the subunit structure as well as the redox-active centers of their intramolecular electron transport chain. Because of these pros- thetic groups, their absorption spectra show far-reaching simi- larities (Fig. 3). Quinoline 2-oxidoreductase from C. testosteroni 63 is a large protein (360 kDa) consisting of three differently sized subunits (87, 32, 22 kDa) occurring in equimolar amounts.

542 Schach et al. (Eur: J. Biochem. 232)

Fig. 6. Conversion of (3-methyl-)quinoline (1) to (3-methyl-)2-oxo-1,2-dihydroquinoline (2), catalysed by quinoline 2-oxidoreductase, and conversion of (3-methyl-)2-0~0-1,2-dihydroquinoIine ( 2 ) to 5,6-dihydro-5,6-dihydroxy-(3-methyl-)2-oxo-l,2-dihydroquinoline (3), catalysed by 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase. R = (-CH,), -H.

An a2Jlpz)12 subunit composition as known, e.g. for quinoline 2- oxidoreductases (Peschke and Lingens, 1991), seems plausible. However, because of the discrepancy between the molecular mass calculated from SDS/PAGE (282 kDa) and the mass ob- served in gel filtration, other possibilities cannot be excluded. The enzyme contains FAD, molybdenum, iron, and acid-labile sulfur in the stoichiometric ratio of 2 : 2: 8 : 8. Liberation of CMP from the purified enzyme indicates that molybdenum is com- plexed to molybdopterin cytosine dinucleotide.

The comparison of the amino-terminal protein sequences of the subunits of quinoline 2-oxidoreductase from C. testosteroni 63 and of corresponding stretches of amino acids of some other molybdenum-containing hydroxylases revealed significant simi- larities (Fig. 4). Especially the N-terminal sequences of the Jl subunits of all three qiiinoline 2-oxidoreductases coincided nearly completely. Very similar to these was the corresponding sequence of quinoline-4-carboxylic acid 2-oxidoreductase from Agrobacterium spec. 1B. Moreover, significant similarities were found between the N-termini of the P-subunits of quinaldine 4- oxidoreductase from Arthrobacter spec. Rii61 a, and of quinaldic acid 4-oxidoreductase from Pseudomonas sp. AK-2. Con- sidering the N-termini of this subunit, all enzymes mentioned seem to be related, but the oxidoreductases hydroxylating at C2 and C4 seem to be closer related to each other.

Whereas the N-terminal sequences of the quinoline 2-oxido- reductases from Pseudomonas putida 86 and from Rhodococcus spec. B1 show complete identity for all three subunits (Fig. 4, sequence 2), the a- and the y-subunit of quinoline 2-oxidoreduc- tase from C. testosteroni 63 differ more. However, for further comparative analysis, the total sequences need to be known be- cause highly conserved regions, such as cofactor-binding do- mains, and varkble regions generally exist.

2-0~0-1,2-dihydroquinoline 5,6-dioxygenase, an aromatic ring dioxygenase, catalyses the second step in (3-methyl-)quino- line degradation (Fig. 6). Dioxygenation of aromatic rings to their cis-dihydrodiols is a common mode in procaryotes to pre- pare these substrates for ring fission. Nevertheless, in the degra- dation of N-heterocyclic compounds, the only dihydrodiol that has been detected as metabolite up to now is the 7,8-dihydrodiol of kynurenic acid in the degradation pathway of kynurenic acid (Kuno et al., 1961 ; Taniuchi and Hayaishi, 1963). Several pieces of evidences indicated the formation of a dihydrodiol in (3- methyl-)quinoline degradation by C. festosteroni 63. The fact that the product of the enzymic conversion of 2-0x0-1 ,2-dihy- droquinoline is optically active reveals its non-aromatic charac- ter. After acidification, this metabolite disappeared, concomitant with formation of 6-hydroxy-2-0x0-1 ,2-dihydroquinoline. It is known that dihydrodiols dehydrate spontaneously upon treat- ment with heat or acid and predominantly yield only one of the two possible monohydroxy derivatives (Taniuchi and Hayaishi, 1963; Fetzner et al., 1993). This would account for the exclusive appearance of 6-hydroxy-2-oxo-l,2-dihydroquinoline. Spraying of TLC plates with two reagents for the detection of vicinal diols led to the same result: the neutral samples gave positive reac- tions, indicating a dihydrodiol structure, whereas acidified sam-

ples did not show any reaction. Since the degradation pathway had been shown to involve 5,6-dihydroxy-3-methyl-2-oxo-1,2- dihydroquinoline as intermediate (Schach et al., 1993), all these results establish 5,6-dihydro-S,6-dihydroxy-(3-methyl-)2-oxo- 1,2-dihydroquinoline as the second metabolite in (3-methyl-)- quinoline degradation by C. testosteroni 63.

Aromatic ring dioxygenases usually are multicomponent en- zyme systems which consist of one or two redox proteins, func- tioning as electron-transport components, and a terminal dioxy- genase, acting as hydroxylase component (Mason and Cam- mack, 1992; Harayama et al., 1992). In the case of 2-0x0-1,2- dihydroquinoline .5,6-dioxygenase, we did not observe a separa- tion into several components with any of the separation methods tried. In contrast, multicomponent oxygenases were reported to completely separate into their components in ion-exchange chro- matography (Yeh et al., 1977; Axcell and Geary, 1975; Ensley et al., 1982; Markus et al., 1986; Fetzner et al., 1989; Romanov and Hausinger, 1994) or gel filtration (Ribbons, 1971 ; Markus et al., 1984). The assumption that 2-0xo-l,2-dihydroquinoline 5,6-dioxygenase is a single-component dioxygenase is further supported by the finding that its specific activity is independent of protein concentration.

To our knowledge, the only other aromatic ring dioxygenase reported to be a single-component enzyme is the naphthalene oxygenase from Corynebacterium renale (Dua and Meera, 1981). This 99-kDa enzyme, which converts its substrate to the corresponding cis-l,2-dihydrodiol, consists of two non-identical subunits.

Because of its instability, we only succeeded in partially pu- rifying 2-0xo-l,2-dihydroquinoline 5,6-dioxygenase. Therefore, some of its properties still remain obscure. However, indirect conclusions on cofactor composition were drawn from inhibitor studies.

2-Oxo-1,2-dihydroquinoline 5,6-dioxygenase was affected by the flavin analogues acriflavin and quinacnne. The participa- tion of a flavin in the electron transfer from NADH to the sub- strate seems probable, since flavin serves as reservoir for electrons and is capable of converting a single two-electron transfer into two one-electron transfers. In the aromatic ring di- oxygenases, this function is fulfilled by one of the redox pro- teins, the (iron-sulfur) flavoprotein with NAD(P)H reductase ac- tivity.

2-Oxo-1,2-dihydroquinoline 5,6-dioxygenase is strictly de- pendent on NADH. With NADPH as electron donor, no activity was measured. The enzyme is activated by exogenously added ferrous iron. A similar effect is known for many aromatic ring dioxygenases, for example toluene dioxygenase (Yeh et al., 1977), benzene dioxygenase (Axcell and Geary, 1975; Geary et al., 1984), and 4-chlorophenylacetate 3,4-dioxygenase (Markus et al., 1986; Schweizer et al., 1987). In multicomponent oxygen- ases, the dioxygen binding is thought to be mediated by a mono- nuclear iron associated with the terminal oxygenase component. This Fe2+ cofactor may be lost during the purification procedure, and replacement of the lost mononuclear iron by exogenously supplied Fe2+ ions is a possible explanation for its activating

Schach et al. (Eui: J. Biochem. 232) 543

effect. Two further results reveal the involvement of iron ions in the reaction mechanism. 2-0x0-I ,2-dihydroquinoline 5,6-dioxy- genase is strongly inhibited by all the chelating agents tested, implying the importance of a loosely bound metal ion in the enzymic reaction. The finding that full enzymic activity is not achieved when Fez+ is added together with Fe3+ can be ex- plained by the competition of both ions for one and the same binding site. The inhibition by sulfhydryl active agents as well as the stabilizing effects by dithiothreitol and (SH),Ery-ol indi- cates the importance of SH groups for catalytic activity of 2- oxo-l,2-dihydroquinoline 5,6-dioxygenase.

Substrate also was a stabilizer for 2-oxo-1,2-dihydroquino- line 5,6-dioxygenase. This effect has been described for 3-hy- droxyphenylacetate 6-hydroxylase (Van Berkel and Van den Tweel, 1991) and for phenol hydroxylase (Neujahr, 1982). In both cases, the protection was attributed to the fact that the bind- ing of substrate intensifies the binding of FAD. Whether this applies for 2-0x0-I ,2-dihydroquinoline 5,6-dioxygenase still re- mains unclear.

The degradation of 3-methylquinoline by C. testosteroni 63 had been reported to proceed via 3-methyl-2-0x0-I ,2-dihydro- quinoline, 5,6-dihydroxy-3-methyl-2-oxo-1,2-dihydroquinoline and 2,5,6-trihydroxy-3-methylpyridine (Schach et al., 1993). With our enzymic studies we have eIucidated the mode of 5,6- dihydroxy-(3-methyl-)2-oxo-l,2-dihydroquinoline formation in C. testosteroni 63. This compound is formed by a dioxygenation and a following dehydrogenation step, not by two subsequent monooxygenation steps. Our results indicated that the conver- sion of (3-methyl-)2-oxo-l,2-dihydroquinoline to the corre- sponding 5,6-dihydrodiol is catalyzed by a single-component aromatic ring dioxygenase. Further studies on this novel enzyme will be interesting to gain more insight into the mechanism of this dioxygenation reaction.

This work was supported by Riitgerswerke AG (Castrop-Rauxel, Germany), by the Fonds der Chemischen Industrie, and by the Deutsche Forschun~sgemeinscha~. We thank Prof. Dr Schreiber (Institut fur Phy- sik der Universitat Hohenheim) for X-ray fluorescence analyses, and Prof. Jung, Dr Stefanovic, and V. Gnau (Institut fur Organische Chemie der Universitat Tubingen) for NH2-terminal amino acid sequence analy- ses.

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