JOURNAL THE BIOLOGICAL CHEMISTRY OF Vol. No. … with concomitant reduction of a suitable elec- tron...

THE JOURNAL OF BIOLOGICAL CHEMISTRY 0 1994 by The American Society for Biochemistry and Molecular Biology, Inc

Vol. 269, No. 15, Issue of April 15, pp. 11254-11260, 1994 Printed in U.S.A.

Purification and Characterization of Isoquinoline 1-Oxidoreductase from Pseudomonas diminuta 7, a Novel Molybdenum-containing Hydroxylase*

(Received for publication, September 20, 1993, and in revised form, January 13, 1994)

Martin Lehmann, Barbara Tshisuaka, Susanne Fetzner, Petra Roger, and Franz LingensS From the Institut fur Mikrobiologie (250), Universitat Hohenheim, 0-70593 Stuttgart, Germany

Isoquinoline 1-oxidoreductase, which catalyzes the hydroxylation of isoquinoline to l-oxo-1,2-dihydroisoquinoline with concomitant reduction of a suitable electron acceptor, was purified from the isoquinoline de- grading bacterium Pseudomonas dirninuta 7 to apparent homogeneity. The native enzyme was a heterodimer with a molecular mass of 96 kDa consisting of a 16- and a 80-kDa subunit. It contained 0.85 g atom molybdenum, 3.95 g atom iron, 3.9 g atom acid-labile sulfur, 2.1 mol of phosphate, and 1 mol of CMP/mol of enzyme. CMP and phosphate are suggested to originate from molybdopterin cytosine dinucleotide of the pterin molybdenum cofactor. It is assumed that the iron and the acid-labile sulfur are arranged in two (2Fe-2s) clusters. The isoelectric point of the isoquinoline 1-oxidoreductase was within the range of pH 6.2 to 6.8. Cytochrome c, ferricyanide, and several non-physiological electron acceptors served as oxidizing substrates, whereas 0, and NAD were not used. Isoquinoline 1-oxidoreductase revealed a high specificity toward the reducing substrates isoquinoline, 5-hydroxyisoquinoline, quinazoline, and phthalazine. Isoquinoline 1-oxidoreductase was inactivated by methanol, arsenite, p-hydroxymercuribenzoate, 1,lO-phenanthroline, and cyanide. Additionally, the enzyme was inactivated upon incubation with its substrates isoquinoline, which slowly inhibited the enzyme in the absence of an electron acceptor, and 5-hydroxyisoquinoline, which rapidly and very effectively inactivated the enzyme in the presence as well as in the absence of the electron acceptors iodonitrotetrazolium chloride, phenazine methosulfate, or ferricyanide.

Contrary to the numerous studies on the catabolism of quinoline (1-71, only a few investigations have been published so far concerning the aerobic, microbial degradation of isoquinoline. Aislabie et al. (8) isolated a n isoquinoline-utilizing Acineto- bacter strain from an enrichment culture. Furthermore, Roger et al. (9) isolated a Pseudomonas diminuta strain and an Al- caligenes faecalis strain capable of utilizat.ion of isoquinoline as sole carbon source. In both cases the authors observed the excretion of l-oxo-l,2-dihydroisoquinoline into the medium (8, 9). Further metabolites have not been detected nor have any studies on the enzymes involved in the breakdown of isoquinoline been carried out so far.

* This work was supported by the Ruetgerswerke AG, Castrop- Rawel, Germany, the Fonds der Chemischen Industrie, and the Deut- sche Forschungsgemeinschaft. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “aduertzsement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

$ To whom correspondence should be addressed. Tel.: 711-459-2222; Fax: 711-459-2238.

In the degradation of related compounds, the initial step is often a nucleophilic attack of the N-heterocycle. For example, nicotinic acid (10, l l ) , picolinic acid (12), nicotine (13, 14), quinoline (1-71, and other quinoline derivatives (15-17) are hydroxylated adjacent to the N-heteroatom, whereas in quinaldine (18) and quinaldic acid (19, 20), the hydroxyl group is incorporated at C-4 of the heterocycle. Probably, all these reac- tions are catalyzed by molybdenum-containing hydroxylases

Among the enzymes catalyzing the hydroxylation of N-het- erocyclic compounds, quinoline 2-oxidoreductases (28, 291, quinoline 4-carboxylic acid 2-oxidoreductase (311, quinaldine 4-oxidoreductase (32), and quinaldic acid 4-oxidoreductase from a Pseudomonas strain (20) appear to be similar to one another. Apart from quinaldic acid 4-oxidoreductase from Ser- ratia marcescens 2CC-1, which is the sole enzyme of this group that consists of two different subunits (cup) and seemingly contains no FAD (19), they all show a cu,p,y, structure and contain eight FeIS, two FAD, and two molybdopterin cytosine dinucleotides. Surprisingly, these molybdenum-containing hydroxylases strongly resemble the CO dehydrogenases from various carboxydotrophic bacteria, especially in subunit structure and cofactor composition (33-35). Apart from the enzymes hydroxy- lating quinoline and its derivatives, molybdopterin cytosine dinucleotide has only been found in the CO dehydrogenases (36, 37).

Up to now, the only molybdenum-containing hydroxylases known to catalyze the conversion of isoquinoline at measurable rates are aldehyde oxidase from rabbit liver (381, which hy- droxylates at C-2 of isoquinoline, and quinaldine 4-oxidoreductase from Arthrobacter spec. Rii 61a (32).

In order to investigate which type of enzyme initiates the degradation of isoquinoline, isoquinoline 1-oxidoreductase was purified from €? diminuta 7. The enzyme was characterized and compared with other molybdenum-containing hydroxylases.

(19-32).

EXPERIMENTAL PROCEDURES Materials-Butyl-Sepharose CL-4B, HiLoad 16/10 S-Sepharose HP,

HiLoad 16/60 Superdex 200 prep grade, low molecular mass standard for SDS-polyacrylamide gel electrophoresis, and PI standards for analytical isoelectric focusing were from Pharmacia Biotech GmbH, Freiburg, Germany. Biolyte 5-7 was from Bio-Rad, Munchen. INT’ was from Serva, Heidelberg, Germany. Precoated Silica Gel Plates SILG- W,,, for thin layer chromatography were from Macherey & Nagel, Duren, Germany. Isoquinoline, quinoline, and quinaldine were a gift from Ruetgerswerke AG, Castrop-Rauxel, Germany. 3-Hydroxybutyric acid was from Fluka AG, Buchs, Switzerland. All other chemicals and biochemicals were obtained from commercial sources and were of the highest purity available.

’ The abbreviations used are: INT, 2-(4-iodophenyl)-3-(4-nitrophenyl)-

methylsulfate; HPLC, high performance liquid chromatography; MCD, 5-phenyl-W-tetrazolium chloride; PMS, N-methyldibenzylpyrazine

molybdopterin cytosine dinucleotide.

11254

Isoquinoline 1-Oxidoreductase 11255

Microorganism and Culture Conditions-P. diminuta 7 was isolated from soil and classified by Roger et al. (9). Strain 7 was grown at 30 "C on mineral salts medium containing(gAiter1: 2.5 SHPO,, 0.85 KH,PO,, 1.0 NaCI, 1.0 (NH,j,SO,, 0.3 MgSO, x 7H,O, 0.03 FeSO, x 7H,O, 0.01 CaCl, x 2H,O, 0.01 Na,MoO, x 2H,O, and vitamin solution (9): 1.0 mMiter. 2 mM isoquinoline was added to the mineral salts medium. In order to determine the influence of tungstate on the degradation of isoquinoline, molybdate was replaced by the same amount of tungstate.

A 100-liter fermenter was used when large amounts of cells were required. The mineral salts medium was supplemented with 2 m isoquinoline and with 4 mM 3-hydroxybutyric acid. Growth was monitored by measuring OD,,, nm. The concentrations of isoquinoline and l-oxo- 1,2-dihydroisoquinoline were estimated by UVNis spectroscopy. After total consumption of isoquinoline and l-oxo-1,2-dihydroisoquinoline, bacteria were fed with 2 mM isoquinoline. 2 h before harvesting the cells by centrifugation 1 nm isoquinoline was added again. The cells were stored at -20 "C.

Enzyme Assays-The isoquinoline 1-oxidoreductase activity was determined by formation of INT-formazan at 503 nm "m = 19.3 m"' cm" (39)) in an optimized assay mixture consisting of 250 pl of 100 mM Tris-HC1 buffer, pH 8.5, with 1.5% Triton X-100, 750 p1 of 5 mM INT solution, 50 pl of 10 mM isoquinoline dissolved in isopropyl alcohol, and 10 pl of enzyme solution. Unless stated otherwise, the reaction was started by addition of substrate. One unit of enzyme activity was de- fined as the amount of enzyme that formed l pmol of formazadmin at 25 "C. Protein estimations were performed according to Lowry et al. (40) with bovine serum albumin as a standard. Enzyme concentration in purified preparations were measured using the following extinction coefficients: E~~~~~ = 152 x lo3 M" cm", E~~~~~ = 35.1 x lo3 M" cm", = 21.3 x lo3 M-' cm", E~~~~~ = 11.7 x lo3 M-I cm".

Isoquinoline 1-oxidoreductase was examined for its substrate specificity with the standard assay except that isoquinoline was replaced by the putative substrates, dissolved in a concentration of 5 mM in isopropyl alcohol or in 50 mM Tris-HC1 buffer, pH 7.5. The products formed were compared by thin layer chromatography with the corresponding hydroxylated compounds. The chromatograms were developed with toluene/dioxane/acetic acid (72:16:1.6) and toluene/ethyl acetate/acetic acid (60:48:8j (6). In order to determine the electron acceptors used by isoquinoline 1-oxidoreductase, INT in the standard assay mixture was replaced by a variety of natural and artificial electron acceptors. The final concentrations (mM) and extinction coefficients used (m"' cm") were as follows: 1,2-benzoquinone 1.89, E~~~~~ = 5.84; nitroblue tetrazolium chloride 1.89, E~~~~ = 15.5; ferricyanide 1.18, E,~,~,,, = 1.02 (41); meldola's blue 0.05, E~~~~~ = 13.9; 2,6-dichlorophenol indophenol 0.01, E~~~~~ = 21 (42); cytochrome c 0.02, E~~~~~ = 21 (43); thionine 0.01, E~~~~~

= 45.0; PMS 0.01, E~~~~~ = 22.0 (44); PMS 0.01 and INT 3.50; benzyl viologen 0.47; methyl viologen 0.94; KNO, 0.94; menadione 3.54; FAD 0.09; FMN 0.19; NAD 0.94; NADP 0.47; methylene blue 0.01; 1.2-naphthoquinone 0.18, and anthraquinone 0.09. Consumption of 0, (in the standard assay without INT) was measured with a Clark-type oxygen electrode (YSI4004, Yellow Springs Instrument Co., Yellow Springs, OH).

The pH optimum was determined using the standard assay replacing Tris-HC1 buffer by Britton-Robinson buffer (45) in the pH range 4-10.

To test putative inhibitors for their time- and concentration-dependent influence on the enzymatic activity, they were incubated together with the isoquinoline 1-oxidoreductase in 20 mM Tris-HC1 buffer, 0.3% Triton X-100, pH 8.5, at room temperature. Aliquots were taken at appropriate intervals for the standard enzyme assay. The effectors tested were as follows: p-hydroxymercuribenzoate in concentrations of 2.5,3.5,5, and 25 p~ with 1.0 unit/ml enzyme; sodium-meta-arsenite (1, 2, 5, and 10 mM with 0.3 unitlml enzyme); potassium cyanide (10, 30, and 50 nm with 0.6 unitlml enzyme); isoquinoline (0.5 mM with 1.6 unitlml enzyme), 5-hydroxyisoquinoline (1.4 and 2.6 PM with 0.42 unitlml enzyme). The conversion of 5-hydroxyisoquinoline was also as- sayed in the presence of 0.06 mM PMS or 0.5, 1,2, and 3.6 mM ferricyanide instead of INT. For methanol inhibition, methanol (final concentration 1.11 M) was added to the standard assay mixture using 1 unitlml enzyme. Quinacrine (0.86 mM final concentration), acriflavine (0.43 mM), EDTA (4.31 mM), 2,2'-bipyridyl (0.43 mM), 1,lO-phenanthroline (0.43 mM), iodoacetate (0.43 m), and various divalent metal ions (0.43 mM final concentration) were also tested for their influence on the enzymatic activity (1.2 uniffml) using the standard test.

Enzyme Purification-All purification steps, unless stated otherwise, were carried out at room temperature. Frozen cells (50 g wet weight) were thawed in 100 ml of 100 m Tris-HC1 buffer, pH 8.0, containing 1 mM EDTA, and disrupted 20 min in a sonifier (model 450, Branson Inc., Danbury, CT) with maximal power. The suspension was centrifuged at

48,000 x g for 40 min at 4 "C, and then the crude extract was incubated at 55 "C for 10 min. After another centrifugation step, (NH,),SO, was added to the supernatant to the final concentration of 0.8 M. Then the (NH,),SO,-enriched protein solution was applied to a butyl-Sepharose CL-4B column (2.5 x 9 cm), which was equilibrated in 50 mM Tris-HC1 buffer, pH 8.0, containing 0.8 M (NH,),SO,. The column was washed with 400 ml of buffer, and the proteins were eluted with a linear gradient (700 ml) of 0.8-0.075 M (NH,),SO, at a flow rate of 0.8 m!h. Active fractions were pooled and dialyzed against 10 mM sodium acetate buffer, pH 5.6, and then concentrated to less than 30 ml by ultrafiltration with a Y"30 membrane (Amicon, Lexington, MA). The concentrated protein solution was applied to S-Sepharose equilibrated in 10 mM sodium acetate buffer, pH 5.6. The column was washed with 50 ml of buffer, and a linear gradient (150 mlj of 0-0.3 M NaCl was applied at a flow rate of 3 mumin. Active fractions were pooled and concentrated to less than 1 ml by ultrafiltration. For gel filtration, the protein solution was loaded in portions of 0.5 ml onto Superdex 200. Proteins were eluted with 50 mM Tris-HC1 buffer, pH 8.0, containing 0.25 M NaC1, a t a flow rate of 1 mumin. Fractions exhibiting enzymatic activity were combined, concentrated by ultrafiltration, and stored a t -80 "C.

Gel Electrophoresis-In order to analyze the homogeneity of isoquinoline 1-oxidoreductase preparations, SDS-polyacrylamide gel electrophoresis was performed using 10% separating and 4% stacking gels (46). Non-denaturing gels were prepared according to Hames (47) using 5% separating and 4% stacking gels in the high pH system. For activity staining, the 1-mm slab gels were immersed in the standard assay mixture. Then the gels were stained in 0.2% (w/v) Coomassie Blue R-250 in water/methanoVacetic acid (40:50:10). Analytical isoelectric focusing was performed in rehydrated gels as recommended by Phar- macia (48). Carrier ampholytes in the range of pH 3-10 and 5.7-8.2 were used. Gels with a higher viscosity were made by addition of 1.5% Triton X-100 or 8 M urea + 1.5% Triton X-100, or 20% glycerol.

Cofactor, Metal, Phosphate, and Acid-labile Sulfur Determinatzons- Two methods were tried to extract FAD from isoquinoline l-oxidoreductase. 1) The enzyme was treated with trichloroacetic acid as described by Meyer (33). 2) Purified enzyme was boiled for 15 min, centrifuged, and the supernatant was examined by UVNis spectroscopy and by HPLC on a LiChrospher RP-8 column (250 x 4.6 mm, 5-pm particle size) according to Buder and Fuchs (49).

Examinations of the nucleotide content of the isoquinoline l-oxidoreductase were performed as described by Frunzke and Meyer (37) and Hettrich et al. (50).

Quantitation of the phosphate content was carried out according to the procedure described by Ames (51). Determination of acid-labile sulfur was performed as described by Beinert (52) using milk xanthine oxidase as a standard.

Multielement analysis of metals like iron and molybdenum was done with a x-ray fluorescence spectrometer (System 77 Finnigan Int. Inc., Sunnyvale, CAI.

Absorption Spectra-Absorption spectra were measured in cells of 1-cm path length a t 25 "C with an Uvicon 930 spectrometer (Kontron Instruments, Neufahrn, Germany).

Molecular Mass Estimation-The native molecular mass of the enzyme was determined by gel filtration on Superdex 200 in 100 mM Tris-HC1 buffer, pH 8.5. Proteins used for calibration were as follows: y-globulin (169 kDa), phosphorylase b (rabbit muscle, 94 kDa), alcohol dehydrogenase (horse liver, 80 kDa), serum albumin (bovine, 67 kDa), albumin (chicken egg, 43 kDa), ribonuclease A (bovine pancreas, 13.7 kDa). The molecular mass of the subunits was estimated using SDS- polyacrylamide gel electrophoresis.

Sedimentation Coeficient-The sedimentation coefficient of purified isoquinoline 1-oxidoreductase was determined in 25 mM Tris-HC1 buffer, pH 8.0, using a BeckmadSpinco model E analytical ultracentrifuge with a photoelectric scanner (Beckman Instruments, Fullerton). The calculation of the sedimentation coefficient and the estimation of the molecular mass of isoquinoline 1-oxidoreductase was carried out as described by Chervenka (53).

RESULTS

Influence of Molybdate and nngs ta te on Zsoquinol ine Degradation-Supplementation of growth medium with high amoun t s of tungstate has been reported to result i n the forma- t ion of inactive demolybdo or tungsten-containing molybdoen- zymes (54-57). Growth of €? dirninuta 7 on isoquinol ine was strikingly diminished in the presence of tungstate, whereas the growth on 1-oxo-1,Z-dihydroisoquinoline was not affected by

11256 Isoquinoline 1-Oxidoreductase TABLE I

Purification of isoquinoline 1-oxidoreductase from I? diminuta 7

Purification step Protein Activity Specific

activity Purification Yield

mg units unitslmg -fold % Crude extract 2185 90 0.04 1 100 Heat treatment 870 85 0.10 2.4 94 Butyl 62 85 1.35 33 93 Sepharose S-Sepharose 7 66 9.64 235 73 Superdex 200 3 46 13.87 338 51

tungstate. These results indicate that the first enzyme of the isoquinoline catabolism, isoquinoline 1-oxidoreductase, is the only molybdenum-containing enzyme in the degradation path- way.

Purification of Zsoquinoline 1-Oxidoreductase-The enzyme was purified 330-fold by heat treatment (55 "C, 10 min), hydro- phobic interaction chromatography on butyl-Sepharose, cation exchange chromatography on S-Sepharose, and a final gel filtration step in a yield of 50%. Table I summarizes the results of a typical purification procedure. The preparation was homoge- neous as shown by native polyacrylamide gel electrophoresis with subsequent activity and protein staining and by analytical ultracentrifugation (data not shown).

Molecular Masses and Subunit Composition-The native molecular mass of purified isoquinoline 1-oxidoreductase was determined to 95 kDa by gel filtration. Analytical ultracentrifugation revealed a sedimentation coefficient of = 5.75 S indicating a native molecular mass of 93 kDa. SDS-polyacrylamide gel electrophoresis revealed that the enzyme is composed of two different subunits with molecular masses of 16 and 80 kDa (Fig. 1). Densitometric scans and integration of Coomassie Blue-stained gels showed a molecular ratio of the subunits of 1.2:l. Thus, we propose that the isoquinoline l-oxidoreductase from I! diminuta 7 exists as a heterodimer (ap).

Isoelectric Focusing-Determination of the isoelectric point was carried out using gels of different viscosity. However, in all cases the enzyme separated into eight distinct bands in a pH range from 6.2 to 6.8. All bands exhibited isoquinoline l-oxidoreductase activity as shown by activity staining. The inten- sity of the individual bands varied from purification to purification. The elution profile of the cation exchange chromatography also indicated that the isoquinoline l-oxidoreductase preparations consisted of fractions with different isoelectric points (data not shown).

Content of Metals, Phosphate, and Acid-labile Sulfur-X-ray fluorescence spectroscopy revealed an average value of 0.85 g atom molybdenum, 3.95 g atom iron, and 0.5 g atom copper/mol of enzyme (based on a molecular mass of the enzyme of 95 kDa). An average value of 3.9 g atom acid-labile sulfur/molecule of enzyme was determined. The absorption spectrum (Fig. 2) indicated that the iron is not bound to a heme-like structure but in iron-sulfur clusters. The phosphate content was 2.1 mol of phosphatdmol of enzyme.

Flavin and Nucleotide Content-Neither heat treatment nor denaturation with trichloroacetic acid extracted a flavin cofactor from isoquinoline 1-oxidoreductase, as shown spectropho- tometrically and by HPLC. One mol of isoquinoline l-oxidoreductase contained 1.0 mol of CMP, while other nucleotides were not detected. The absorption spectrum, the failure to extract a flavin, and the absence ofAMP, which would derive from FAD hydrolysis, show that the enzyme contains no FAD. CMP is generated by hydrolysis from the MCD molybdenum cofactor. Therefore, the 2.1 mol of phosphate foundmol of enzyme de- rived from molybdopterin cytosine dinucleotide.

Absorption Spectra-The UVNis spectrum of the red-brown isoquinoline 1-oxidoreductase as isolated (Fig. 2) had a protein

A B 1-oxidoreductase. Lane A, molecular mass standards (in kDa): phos-

FIG. 1. SDS-polyacrylamide gel electrophoresis of isoquinoline

phorylase h, 94; bovine serum albumin, 67; ovalbumin, 43; carbonic anhydrase, 30; soybean trypsin inhibitor, 20.1; lactalbumin, 14.4. Lane B, purified isoquinoline 1-oxidoreductase (see "Experimental Proce- dures").

0 0 C 0

g 1.0 -

v) .n 4 400 500 600

0.5 - Wavelength [nrn]

0 . 0 " " " " " " ' " " " " ' 300 400 500 600 700

Wavelength [nm]

FIG. 2. Absorption spectra of isoquinoline 1-oxidoreductase from P. diminuta. 1.25 mg/ml enzyme was dissolved in 50 mM Tris- HCI buffer, pH 8.0. Inset, solid line, enzyme as isolated; dash line; enzyme reduced with 79 PM isoquinoline; dotted line, difference spectrum between spectrum of enzyme as isolated and spectrum of substrate-reduced enzyme.

maximum at 280 nm, a broad double peak at 429 and 469 nm, and two shoulders in the regions of 338 and 550 nm. The A,,d A,,, ratio of the enzyme was about 8.0, and the A,,dA,,, ratio was 2.0. The maxima at 429 and 469 nm, and the shoulder at 550 nm disappeared upon treatment of the enzyme with isoquinoline or with sodium dithionite (Fig. 2). In the range of 400 to 600 nm, the difference spectrum had a maximum at 472 nm


a "&* & Acceptor Acceptor x 2 H

quinoline catalyzed by isoquinoline 1-oxidoreductase. FIG. 3. Conversion of isoquinoline to l-oxo-1,2-dihydroiso-

= 9040 M - ~ cm") and shoulders at 430 and 550 nm. Reduction of the enzyme with isoquinoline or with sodium dithionite yielded identical absorption spectra.

Enzyme Assay-The isoquinoline 1-oxidoreductase catalyzes the hydroxylation of isoquinoline to l-oxo-1,2-dihydroisoquinoline with concomitant reduction of INT (Fig. 3). From the highest specific activity of 15 unitslmg, a turnover number of 1455 min" and a corresponding turnover time of 4.1.10-' s was calculated. Table I1 shows the relative isoquinoline l-oxidoreductase activity with various electron acceptors based on the activity with INT in the enzyme standard assay. No activity was measured using the electron acceptors methyl viologen, benzyl viologen, KNO,, menadione, methylene blue, FAD and FMN, NAD and NADP, 1,2-naphthoquinone, anthraquinone, and 0,.

The pH optimum of isoquinoline 1-oxidoreductase was -8.5. The optimum temperature for the enzyme reaction was in the range of 55 to 60 "C. This was consistent with examinations of the temperature stability of the enzyme (incubated for 5 min in 50 mM Tris-HC1 buffer, pH 8.01, which decreased rapidly above 60 "C.

Substrate Specificity-A wide variety of quinoline and pyridine derivatives was tested in the standard assay. The enzyme hydroxylated isoquinoline (100% activity) to 1-oxo-1,2- dihydroisoquinoline, quinazoline (45%) to 4-hydroxyquinazo- line, 5-hydroxyisoquinoline to 1,5-dihydroxyisoquinoline, and phthalazine (89%). 5-Hydroxyisoquinoline very effectively inactivated the enzyme (see below). The following compounds were not accepted as substrates: quinoline, 2-/4-/8-monochloro- quinoline, 2-/3-/4-monomethylquinoline, quinoline 2-13-14-15-16- /7-/8-monocarboxylic acid, 2-14-16-17-/8-monohydroxyquinoline, 2,4-dihydroxyquinoline, 8-chloroquinaldine, l-methylisoquino- line, 1,5-dihydroxyisoquinoline, l-oxo-1,2-dihydroisoquinoline, xanthine, hypoxanthine, allopurinol, cinnoline, quinoxaline, adenine, guanine, naphthalene, pyridine, pyridine 2-carboxylic acid, nicotinic acid, kynurenic acid, pyridine 2,3-dicarboxylic acid, and formamide.

The K, values for isoquinoline and INT, determined in the standard assay and calculated in Lineweaver-Burk plots, were 1.6 . 10 -5 M and 1.34 . M, respectively. Inhibitors-p-Hydroxymercuribenzoate, cyanide, and arsen-

ite were tested for their influence on isoquinoline l-oxidoreductase. In each case, a time-dependent decrease in the enzymatic activity was observed, which was markedly dependent upon the concentration of the inhibitor (Fig. 4 and Table 111). In all three cases, the inhibition did not proceed to complete inactivation of the enzyme but to an inhibited steady state, which was also dependent upon the inhibitor concentration used. Methanol inhibited the enzyme only during turnover of isoquinoline. Isoquinoline 1-oxidoreductase was also inactivated upon aerobic incubation with its substrates isoquinoline and 5-hydroxyisoquinoline (Fig. 5). Aerobic incubation of enzyme together with 5-hydroxyisoquinoline resulted in a fast inactivation reaction (Table I11 and Fig. 5). The inactivation was slowed down by addition of electron acceptors like INT, PMS, or ferricyanide. However, the acceptors did not prevent the complete inactivation of the enzyme. The inactivation was also drastically slowed down by addition of isoquinoline to the incubation mixture (Fig. 5). Therefore, isoquinoline and 5-hydroxyisoquinoline presumably react at the same site of the

TABLE I1 Electron acceptors utilized by the isoquinoline I-oxidoreductase from

P: diminuta 7

Electron acceptor Relative rate

%

INT" 100 Meldola's blue 315

PMS 112 Nitroblue tetrazolium chloride 94 Cytochrome c 87 Ferricyanide 86 1,2-Benzoquinone 39 2,6-Dichlorophenol indophenol 4

Standard assay.

INT-PMS mix 204

100 v

80 k 3

5 6 0

40

.- L

0 5 10 1 5 2 0 2 5 30 35

Time [rnin]

FIG. 4. Inactivation of isoquinoline 1-oxidoreductase by arsenite, p-hydroxymercuribenzoate, and cyanide. The enzyme was incubated aerobically with either 30 mM cyanide (0, 0.6 unidml enzyme) or 3.5 pMp-hydroxymercuribenzoate (0,l .O unit/ml enzyme) or 0.95 mM arsenite (m, 0.3 unit/ml enzyme). At appropriate time intervals an aliquot was taken, and the residual activity was determined using the enzyme standard assay (see "Experimental Procedures"). The initial activity was determined without inhibitor, and for the calculation of the 100% value, the dilution caused by addition of the inhibitor was taken into account.

TABLE I11 Effect of various inhibitors on the isoquinoline

I-oxidoreductase activity The half-lifes were calculated from plots (see Fig. 4) resulting from

incubation of isoquinoline 1-oxidoreductase in 20 nm Tris-HC1 buffer, pH 8.5, 0.3% Triton X-100, with the various inhibitors. At appropriate time intervals an aliquot was taken, and the activity was determined in the standard assay.

Inhibitor Cone. Half-life

mM nin KCN 50.0

30.0 10.0

2.6 5.9

45.2 Arsenite 9.50 0.8

1.90 0.95 12.9

6.1

p-Hydroxymercuribenzoate 0.0250 0.0050

0.6

0.0025 1.9 5.9

5-Hydroxyisoquinoline 0.0014 0.0026

1.9 1.0

Isoquinoline 0.50 159 5-Hydroxyisoquinoline/Isoquinoline 0.0026/0.26 54

enzyme. Incubation with 1.39 p 5-hydroxyisoquinoline (a molecular ratio of substrate to enzyme -4.7) did not lead to complete inactivation of the isoquinoline 1-oxidoreductase probably due to the total conversion of 5-hydroxyisoquinoline. Enzyme spectra, measured during the inactivation process of isoquinoline 1-oxidoreductase by 5-hydroxyisoquinoline revealed that the FelS centers of the enzyme were first reduced (see spectrum after 1 min, Fig. 6). Between 1 and 5 min the FeIS centers

11258 Isoquinoline 1-Oxidoreductase

100

80

6 0

$ 40

Q 20

- K u

+ .- >

V

A

h. I r

0 10 20 30 40 50

Time [min] B

100

80 - R Y 6 0

$ 40

20

0

L z .-

0 4

0 500 1000 1500

T ime [ rn in ]

FIG. 5. Inactivation of isoquinoline 1-oxidoreductase by its substrates isoquinoline and 5-hydroxyisoquinoline. The enzyme was incubated aerobically with its substrates isoquinoline and 5-hydroxyisoquinoline (see "Experimental Procedures"). A, 0.42 unitfml enzyme was inactivated by either 2.63 VM 5-hydroxyisoquinoline (0, a molecular ratio of substrate to enzyme about 8.9) or 1.39 J ~ M 5-hydroxyisoquinoline (0, a ratio of 4.7) or a mixture of 0.26 mM isoquinoline and 2.63 5-hydroxyisoquinoline (W). B, 1.6 unit/ml enzyme was incubated with either 0.5 nm isoquinoline (0) or a mixture of 0.25 mM isoquinoline and 0.05 mM 5-hydroxyisoquinoline (01, and as references, the enzyme was incubated with the used buffer (W) and with an equivalent concentration of isopropyl alcohol (01, in which the substrates were dissolved.

0.0

1 m i n

-0.1 400 500 600 7 0 0

wave leng th [ nm]

FIG. 6. Difference spectra of 5-hydroxyisoquinoline-treated minus native isoquinoline 1-oxidoreductase. The spectra were re- corded during the aerobic inactivation of 2.8 mg/ml enzyme in 20 mM Tris-HC1 buffer, pH 8.0, by 0.5 mM 5-hydroxyisoquinoline, dissolved in isopropyl alcohol, after 1, 3, 5, 7, and 19 min.

rapidly reoxidized and a significant difference spectrum (e.g. spectrum after 19 min, Fig. 6) with a maximum at 518 nm arose. The product of the conversion of 5-hydroxyisoquinoline by isoquinoline 1-oxidoreductase, 1,5-dihydroxyisoquinoline, affected neither the reduced nor the oxidized form of the enzyme.

The inactivation of isoquinoline 1-oxidoreductase upon aerobic incubation with isoquinoline in the absence of an electron acceptor progressed very slowly in comparison to the inactivation on treatment with 5-hydroxyisoquinoline (Table 111). In contrast to the inactivation of the enzymatic activity by 5-hydroxyisoquinoline, the time course of the inactivation by isoquinoline fitted pseudo-first-order kinetics (Fig. 5).

While 1,lO-phenanthroline decreased the enzymatic activity to -40%, quinacrine, acriflavine, EDTA, 2,2'-bipyridyl, and io- doacetamide did not affect the isoquinoline 1-oxidoreductase activity. Hg2' (84% inhibition), Zn2+ (46%), Mn2+ (40%), Co2+ (39%), and Cu2+ (18%) inhibited the activity of the enzyme, too.

DISCUSSION

From the data of the chemical analyses and the UVNis absorption spectrum of the enzyme, we conclude that the isoquinoline 1-oxidoreductase from I! diminuta 7 is composed of two subunits (80 and 16 kDa) and probably contains two (2Fe- 2s) centers and one MCD molybdenum cofactor as redox active centers. The molybdenum center is of the monooxo-monosul- fido-type, as proved by inactivation of the enzyme with cyanide.

In all molybdenum-containing enzymes apart from the nitro- genase, the metal is coordinated to a pterin molybdenum cofactor. One form, molybdopterin, is found in eucaryotic molyb- doenzymes like milk xanthine oxidase (58) and sufite oxidase from chicken liver (581, in procaryotic xanthine dehydrogenases such as from Pseudomonas putida 86 (50) and from Pseudomo- nas aeruginosa (591, and in some hyperthermophilic archaea (60), where molybdenum is replaced by tungsten. Furthermore, various molybdopterin dinucleotides were discovered as part of the pterin molybdenum cofactors of some procaryotic enzymes. In these dinucleotides, molybdopterin is linked via a pyrophos- phate bond to GMP (61-631, AMP (64), HMP (64), or CMP, which was determined as a component of isoquinoline l-oxidoreductase. Up to now, MCD was only found in CO dehydrogenase from Pseudomonas carboxydoflava (36), quinoline 2-oxidoreductases from Pseudomonas putida 86 and Rhodococcus spec. B1 (50), quinaldic acid 4-oxidoreductase from Pseudomo- nas spec. AK-2 (20), quinaldine 4-oxidoreductase from Ar- throbacter spec. Ru6la (32), and quinoline 4-carboxylic acid 2-oxidoreductase from Agrobacterium spec. B1 (31). However, nothing is known about the reasons for the distribution of the variants of the pterin part among the enzymes containing the pterin molybdenum cofactor.

The absorption spectrum of isoquinoline 1-oxidoreductase lacks the typical flavin maximum at 450 nm of the molybdo- irodsulfur flavoproteins like xanthine or aldehyde oxidase (65). However, the absorption spectrum resembles the spectra of the molybdo-irodsulfur proteins quinaldic acid 4-oxidoreductase from Serratia marcescens 2CC-1 (19) and aldehyde oxidase from Desulfovibrio gigas (66). Furthermore, the absorption spectrum of isoquinoline 1-oxidoreductase shows considerable similarities to those of the deflavo forms of milk xanthine oxidase (67, 68) and of rabbit liver aldehyde oxidase (69). Because of the absorption spectrum and, since it was not possible to extract flavin from the enzyme and to inhibit the enzyme by quinacrine, we conclude that isoquinoline 1-oxidoreductase is a molybdo-irodsulfur protein. There was no indication that the enzyme might lose a putative flavin during the purification procedure. Therefore, besides the quinaldic acid 4-oxidoreductase from S. marcescens 2CC-1 (19), the isoquinoline l-oxidoreductase from P diminuta 7 is the second enzyme involved in the catabolism of N-heterocycles that does not contain a flavin cofactor.

Isoquinoline 1-oxidoreductase is inactivated by arsenite in a pseudo-first-order reaction followed by an inhibited steady state. The enzyme is not reactivated by a washing procedure. This behavior and the difference spectrum (data not shown) of inactivated isoquinoline 1-oxidoreductase closely resemble the inhibition of milk xanthine oxidase by arsenite (70-72). George and Bray (72) suggested an inhibitor complex for milk xanthine oxidase which is formed by arsenite and the cyanolyzable sulfur of the molybdenum center. They also proposed an analogous complex for the reaction of reduced milk xanthine oxidase with p-hydroxymercuribenzoate (73). In contrast to milk xanthine oxidase, isoquinoline 1-oxidoreductase was inactivated by p - hydroxymercuribenzoate in its oxidized state. However, it was shown that the oxidized state of the aldehyde oxidase from rabbit liver was also inhibited by p-hydroxymercuribenzoate


(74). Considering that the arsenite inhibitor complex was formed both with the oxidized (=SI and the reduced (-SH) state of the molybdenum center of milk xanthine oxidase, it is likely that the oxidized state of isoquinoline 1-oxidoreductase forms an analogous Mo-S-Hg inhibitory complex with p-hydroxymercuribenzoate. The specificity of the p-hydroxymercuribenzoate inactivation was demonstrated by the failure to affect the enzymatic activity by iodoacetate.

Another feature, typical for molybdenum-containing hydroxylases, is the inactivation of isoquinoline 1-oxidoreductase by methanol during substrate turnover. From the characteris- tic MOW) EPR signal of methanol-inhibited milk xanthine oxidase (75) and rabbit liver aldehyde oxidase (76, 77), it was concluded that methanol or more likely formaldehyde, which is probably formed by oxidation of methanol at the molybdenum center, traps the molybdenum in the pentavalent state and thus prevents its further redox cycling.

Isoquinoline 1-oxidoreductase was inactivated upon incubation with isoquinoline in the absence of an electron acceptor in a first-order process. In addition, the enzyme was also inactivated by sodium sulfide and sodium dithionite (data not shown). Considering that isoquinoline 1-oxidoreductase is prac- tically unable to use 0, as electron acceptor, we conclude that the permanent fully reduced state of the enzyme due to substrate or reducing reagents provokes an irreversible alteration of the enzyme. Probably, the alteration is located at the molybdenum center, as reported for the xanthine dehydrogenase from Chlamydomonas rheinhardtii (78). Similar substrate inactiva- tions were reported for the aldehyde oxidase from rabbit liver (79).

5-Hydroxyisoquinoline is interesting as substrate as well as inhibitor of isoquinoline 1-oxidoreductase. There are several indications that only the reduced state of isoquinoline l-oxidoreductase is inactivated by 5-hydroxyisoquinoline, while the oxidized state converts 5-hydroxyisoquinoline to 1,5-dihydroxyisoquinoline. Addition of electron acceptors like PMS, INT, or ferricyanide prolonged the phase of conversion of 5-hydroxyisoquinoline but did not prevent the inactivation. Spectra re- corded during the inactivation process (Fig. 6) revealed that the enzyme was entirely reduced for less than 1 min. The Fe/S centers were then slowly reoxidized by oxygen indicating that the enzyme was not capable to oxidize 5-hydroxyisoquinoline any more and that the Fe/S centers have no importance for the inactivation process. Finally, the typical difference spectrum emerged. The delay of the inactivation process by addition of isoquinoline indicates that the molydenum center is the site of the 5-hydroxyisoquinoline reaction. The resulting difference spectrum (Fig. 6) resembles that of milk xanthine oxidase treated with 4,6-dimercaptopyrazolo-[3,4-d]-pyrimidine (80). This compound is related to allopurinol, a potent “suicide” substrate of xanthine oxidase (80). The product of the enzymatic conversion of allopurinol by xanthine oxidase, alloxanthine, formed a complex with Mo(1V) (80). When 0, as electron acceptor was replaced by ferricyanide or PMS, the formation of the inhibitory complex was prevented, and furthermore, inactivated xanthine oxidase was reactivated upon incubation with PMS or ferricyanide (80). However, neither the oxidized nor the reduced isoquinoline 1-oxidoreductase was affected by 1,5-dihydroxyisoquinoline, and in contrast to the inactivation of xanthine oxidase by allopurinol, incubation with ferricyanide or PMS did not prevent the inactivating effect of 5-hydroxyisoquinoline. We assume that 5-hydroxyisoquinoline reacts at the (partially) reduced molybdenum center and forms a stable complex either with Mo(IV), similar to the alloxanthine- (81) or 8-bromoxanthine-molybdenum complex (82), or with Mo(V), similar to the complex formed by formaldehyde (75, 83).

The recently published quinaldic acid 4-oxidoreductase from

S. rnarcescens 2CC-1 (19) resembles the isoquinoline l-oxidoreductase from r! diminuta 7. The structure of these enzymes is unique among the molybdenum-containing hydroxylases (84). The other known procaryotic molybdenum- containing hydroxylases catalyzing the conversion of quinoline or its derivatives contain two FAD, eight Fe/S (it was shown only for the quinoline 2-oxidoreductase to contain two different Fe/S centers (85)), and two MCD molybdenum cofactors. They are composed of three different subunits in an a,P,y, structure with native molecular masses of 300-340 kDa and subunit molecular masses of 80-85 (a ) , 32-35 ( P ) , and 18-22 kDa ( y ) (20, 28, 29, 31,32). Furthermore, the CO dehydrogenases from various carboxydotrophic bacteria (34, 35) and the nicotinic acid dehydrogenase from Bacillus niacini (24) possess a similar structure. The molecular masses of the subunits of isoquinoline 1-oxidoreductase and the quinaldic acid 4-oxidoreductase from S. rnarcescens 2CC-1(19) correspond to the molecular masses of the large and small subunit of the enzymes with a2P2y2 structure. Intermediate forms between the more simple heterodimeric structure of isoquinoline 1-oxidoreductase and quinaldic acid 4-oxidoreductase and the complex a2P2y2 enzymes are nicotine dehydrogenase from Arthrobacter oxidans (25) and 6-hydroxynicotinic acid dehydrogenase from B. niacini (24), which consist of three different subunits (a&) corresponding to a half-molecule of the a,P,y, enzymes.

It is tentatively assumed that the xanthine dehydrogenases from Drosophila melanogaster (84) and rat liver (861, both eucaryotic molybdenum-containing hydroxylases, which show native molecular masses similar to those of the complex procaryotic enzymes, consist of three different domains corresponding in their molecular masses to the three subunits of the large procaryotic hydroxylases. For both enzymes, it is suggested that the small domain contains the Fe/S centers, the middle one the FAD, and the large domain the pterin molybdenum cofactor (84,861. This cofactor distribution among the domains might correspond to the structure of the procaryotic molybdenum-containing hydroxylases (see also Ref. 34). The middle subunit of the aPy/a,P,y, enzymes might contain the FAD, which is missing in the more simple heterodimeric enzymes, and the other two cofactors are distributed among the remain- ing subunits.

On the one hand, genetic studies will have to be performed to elucidate the relationship between the procaryotic and the eucaryotic molybdenum-containing hydroxylases, and on the other hand, EPR studies will have to be done to throw light upon the structure of the Fe/S centers and the complex formed by 5-hydroxyisoquinoline.

Acknowledgments-We are indebted to Prof. Dr. Schreiber and M. Hepp, Institut fur Physik der Universitat Hohenheim, for recording x-ray fluorescence spectra and to K. Kapassakalis for skillful operation of the fermenter.

REFERENCES

2. Bennett, J. L., Updegraff, D. M., Pereira, W. E., and Rostad, C. E. 119851 1. Grant, D. J. W., and Al-Najjar, T. R (1976) Microbios 15, 177-189

3. Shukla, 0. P. 11986) Appl. Enuiron. Microbiol. 51, 1332-1342 4. Shukla, 0. P. 11987) Bid . Mern. 13, 115-131 5. Shukla, 0. P. 11989) Microbios 59, 4 7 4 3 6. Schwarz, G., Bauder, R., Speer, M., Rommel, T. O. , and Lingens, F. (19891 B i d .

7. Bott. G., and Lingens, F. (1991) Biol. Chem. Hoppe-Seyler 372, 381-383 8. Aislabie, J., Rothenburger, S., and Atlas, R. M. 11989) Appl. Enuiron. Micro-

9. Roger, P., Erben, A,, and Lingens, F. 11990) Bid. Chem. Hoppe-Seyler 371,

Microbios Lett. 29, 147-154

Chem. Hoppe-Seyler 370, 1183-1189

biol. 55, 3247-3249

511-513 10. Behrman, E. J., and Stanier, R. Y. 11957) J. Biol. Chem. 228,923-945 11. Hunt, A. L., Hughes, D. E., and Lowenstein, J. M. (1958) Biochem. J . 69,

12. Dagley, S., and Johnson, P. A. (1963) Biochim. Biophys. Acta 78, 577-587 13. Hochstein, L. I., and Dalton, B. P. (19651 Biochem. Bioph.ys. Res. Cornmun. 21,

170-173

644-648

11260 Isoquinoline 1-Oxidoreductase 14. 15.

16.

17.

18.

19. 20.

22. 21.

23. 24. 25.

26. 27.

28.

29. 30.

31. 32. 33. 34.

Hochstein, L. I . , and Dalton, B. P. (1967) Biochim. Biophys. Acta 139, 56-68 Schmidt, M., Roger, P., and Lingens, F. (1991) Biol. Chem. Hoppe-Seyler 372,

Schach, S., Schwarz, G., Fetzner, S., and Lingens. E (1993) Biol. Chem. Hoppe- 1015-1020

Seyler 374, 175-181

479-488

371. 1005-1008

..

Ruger, A., Schwarz, G., and Lingens, F. (1993) Biol. Chem. Hoppe-Seyler 374,

Hund, H.-K., de Beyer, A., and Lingens, F. (1990) Biol. Chem. Hoppe-Seyler

Fetzner, S., and Lingens, F. (1993) Biol. Chem. Hoppe-Seyler 374, 363-376 Sauter, M., Tshisuaka, B., Fetzner, S., and Lingens, F. (1993) B i d . Chem.

Hirschberg, R., and Ensign, J. C . (1971) J. Bacteriol. 108, 751-756 Dilworth, G. L. (1983)Arch. Biochem. Biophys. 221,565-569 Nagel, M., and Andreesen, J . R. (1989) FEMS Micrpbiol. Lett. 59, 147-152 Nagel, M., and Andreesen, J . R. (1990) Arch. Microbiol. 154, 605413 Freudenberg, W., Koenig, K., and Andreesen, J. R. (1988) FEMS Microbiol.

Tate, R. L., and Ensign, J . C. (1974) Can. J. Microbiol. 20, 695-702 Siegmund, I., Koenig, K., and Andreesen, J. R. (1990) FEMS Microbiol. Lett.

Bauder, R., Tshisuaka, B., and Lingens, F. (1990) Biol. Chem. Hoppe-Seyler

Peschke, B., and Lingens, F. (1991) Biol. Chem. Hoppe-Seyler 372, 1081-1088 Blaschke, M., Kretzer, A., Schafer, C., Nagel, M., and Andreesen, J. R. (1991)

Bauer, G., and Lingens, F. (1992) Biol. Chem. Hoppe-Seyler 373,699-705 de Beyer, A., and Lingens, F. (1993) B i d . Chem. Hoppe-Seyler 374, 101-110 Meyer, 0. (1982) J. Biol. Chem. 257, 1333-1341 Meyer, O., Jacobitz, S., and Kruger, B. (1986) FEMS Microbiol. Reu. 39, 161-

Hoppe-Seyler 374, 1037-1046

Lett. 52, 13-18

67,281-284

371, 1137-1144

Arch. Microbiol. 155, 164-169

1 7 0

35. Morsdorf, G., Frunzke, K., Gadkari, D., and Meyer, 0. (1992) Biodegradation

36. Johnson, J. L., Rajagopalan, K. V.. and Meyer, 0. (1990)Arch. Biochem. Bio-

37. Frunzke, K., and Meyer, 0. (1991) BIOforum 14,83, P 278 38. Stubley, C., Stell, J. G . P., and Mathieson, D. W. (1979)Xenobiotica 9,475484 39. Babson, A. L., and Babson, S. R. (1973) Clin. Chem. 19,766-769 40. Lowry, 0. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J . (1951) J. Biol.

41. Schellenberg, K. A., and Hellerman, L. (1958) J. Biol. Chem. 231, 547-556

43. Massey, V. (1959) Biochim. Biophys. Acta 34,255-256 42. Armstrong, J. M. (1964) Biochim. Biophys. Acta 86, 194197

44. Dixon. M. (1971) Biochim. Biophys. Acta 226, 259-268 45. Britton, H. T. S., and Robinson R. A. (1931) J. Chem. SOC. 1456-1462 46. Schagger, H., and von Jagow, G . (1987) Anal. Biochem. 166,368-379 47. Hames, B. D. (1990) in Gel Electrophoresis of Proteins (Hames, B. D., and

48. Schickle, H., and Westermeier, R., Pharmacia Bulletin SDRE-01, Freiburg,

49. Buder, R., and Fuchs, G. (1989) Eul: J. Biochem. 185,629-635 50. Hettrich, D., Peschke, B., Tshisuaka, B., and Lingens, F. (1991) Biol. Chem.

51. Ames, B. N. (1966) Methods Enrymol. 8, 115-118 52. Beinert, H. (1983) Anal. Biochem. 131,373-378 53. Chervenka, C . H. (1969) A Manual of Methods for the Analytical Ultracentri-

Ll.7

3, 61-82

phys. 283, 542-545

Chem. 193,265-275

Rickwood, D., eds), 2nd Ed., p. 36, IRL Press, Oxford

Germany

Hoppe-Seyler 372,513-517

54.

55.

56.

51. 58.

59.

60.

61.

62. 63.

64. 65. 66.

67. 68.

69. 70.

71.

Johnson, J . L., Rajagopalan, K. V., and Cohen, H. J. (1974) J. Biol. Chem. 249,

Johnson, J. L., Cohen, H. J., and Rajagopalan, K. V. (1974)J. Biol. Chem. 249,

Johnson, J . L., Waud, W. R., Cohen, H. J., andRajagopalan, K. V. (1974)J. Biol.

Johnson, J. L., and Rajagopalan, K. V. (1976) J. Bid. Chem. 251,5505-5511 Kramer, S. P., Johnson, J. L., Ribeiro,A.A., Millington, D. S., and Rajagopalan,

Johnson, J. L., Chaudhury, M., and Rajagopalan, K. V. (1991) BioFactors 3,

Johnson. J . L., Rajagopalan, K. V., Mukund, S., and Adams, M. W. W. (1993) J.

Johnson, J. L., Bastian, N. R., and Rajagopalan, K. V. (1990) Proc. Natl. Acad.

Karrasch, M., Borner, G., and Thauer, R. K. (1990) FEES Lett. 274.48-52 Johnson, J . L., Bastian, N. R., Schauer, N. L., Ferry, J . G., and Rajagopalan, K.

Borner, G., Karrasch, M., and Thauer, R. K. (1991) FEES Lett. 290,31-34 Rajagopalan, K. V., and Handler, P. (1964) J. Bid. Chem. 239, 1509-1514 Moura, J. J. G., Xavier, A. V., Bruschi, M., Le Gall, J., Hall, D. O., and Cam-

Komai, H., Massey, V., and Palmer, G. (1969) J. Biol. Chem. 244, 1692-1700 Kanda, M., Brady, E O., Rajagopalan, K. V., and Handler, P. (1972) J. Biol.

Branzoli, U., and Massey, V. (1974) J. Bid. Chem. 249, 4339-4345 Coughlan, M. P., Rajagopalan, K. V, and Handler, P. (196915. Biol. Chem. 244,

Hille, R., Stewart, R. C., Fee, J. A,, and Massey, V. (1983) J. Bid . Chem. 258,

fuge, pp. 23-33, Beckman Instruments Inc., Palo Alto, CA

859-866

5046-5055

Chem. 249, 5056-5061

K. V (1987) J. Biol. Chem. 262, 16357-16363

103-107

Biol. Chem. 268,4848-4852

Sci. U. S. A. 87, 319LL3194

V (1991) FEMS Microbiol. Lett. 77, 213-216

mack, R. (1976) Biochem. Biophys. Res. Comm. 72, 782-789

Chem. 247, 765-770

2658-2663

72. George, G. N., and Bray, R. C. (1983) Biochemistry 22, 1013-1021 73. George, G. N., and Bray, R. C. (1983) Biochemistry 22,5443-5452 74. Rajagopalan, K. V., Fridovich, I . , and Handler, P. (1962) J. Biol. Chem. 237,

75. Pick, E M., McGartoll, M. A,, and Bray, R. C. (1971) Eul: J . Biochem. 18,65-72 76. Rajagopalan, K. V., Handler, P., Palmer, G., and Beinert, H. (1968) J. Biol.

77. Raiagopalan. K. V.. Handler, P., Palmer, G., and Beinert, H. (1968) J. Bid.

48494856

922-928

Chem. 243, 3784-3796 . " Chem. 243,3797-3806

chim. Biophys. Acta 1117, 159-166 78. Perez-Vicente, R., Alamillo, J . M., Cdrdenas, J., and Pineda, M. (1992) Bio-

79. Coughlan, M. P., Johnson, J . L., and Rajagopalan, K. V. 11980) J. Biol. Chem.

80. Massey, V., Komai, H., Palmer, G., and Elion, G . B. (1970) J. Bid. Chem. 245,

81. Hawkes, T. R., George, G . N., and Bray, R. C. (1984)Biochem. J. 218,961-968 82. Hille, R., and Stewart, R. C. (1984) J. Bid. Chem. 259, 1570-1576 83. Bray, R. C . (1988) Q. Reu. Biophys. 21,299-329 84. Wootton, J. C., Nicolson, R. E., Cock, J. M., Walters, D. E., Burke, J. F., Doyle,

W. A,, and Bray, R. C. (1991) Biochim. Biophys. Acta 1057, 157-185 85. Tshisuaka, B., Kappl, R., Huttermann, J., and Lingens, F. (1993) Biochemistry

255,2694-2699

2837-2844

86. Amaya,Y.,Yamazaki, K., Sato, M., Noda, K., Nishino, T., and Nishino, T. (1990) 32,12928-12934

J. Biol. Chem. 265, 14170-14175

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