©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Role of Cysteine 337 and Cysteine 340 in Flavoprotein That Functions as NADH Oxidase from Amphibacillus xylanus Studied by Site-directed Mutagenesis (*)

(Received for publication, October 12, 1994; and in revised form, December 27, 1994)

Kenji Ohnishi (1) Youichi Niimura (2)(§) Makoto Hidaka (3) Haruhiko Masaki (3) Hideo Suzuki (2) Takeshi Uozumi (3) Takeshi Nishino (4)

From the  (1)Department of Agricultural Chemistry, Tokyo University of Agriculture, Tokyo 156, the (2)Department of Food Science and Technology, Tokyo University of Agriculture, Hokkaido 099-24, the (3)Department of Biotechnology, The University of Tokyo, Tokyo 113, and the (4)Department of Biochemistry and Molecular Biology, Nippon Medical School, Tokyo 113, Japan

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

A flavoprotein from Amphibacillus xylanus catalyzes the reduction of oxygen to hydrogen peroxide. Each polypeptide chain in the tetrameric enzyme contains 5 cysteine residues. The complete reduction of enzyme by dithionite requires 6 electrons. Such behavior indicates the presence of redox centers in addition to the FAD, and these could be disulfides. In order to assess the catalytic role of disulfide in the enzyme, 2 of the cysteines (Cys-337 and Cys-340), which show a high degree of homology with alkyl hydroperoxide reductase F52a protein and thioredoxin reductase, have been changed to serines by site-directed mutagenesis of the cloned flavoprotein gene (individually and in a double mutant). Titration of the three mutant enzymes, lacking Cys-337, Cys-340, or both cysteines, requires only 2 electron eq to reach the reduced flavin state. These results indicate the absence of a redox-active disulfide and demonstrate the involvement of Cys-337 and Cys-340 in the redox-active disulfide. The catalytic activity of the three enzymes was examined by steady-state analysis. The K for NADH and oxygen and the k value of these mutant enzymes were essentially the same as those of wild type. The NADH oxidase activities were also accelerated markedly in the presence of free FAD, which is the case for wild-type enzyme. The NADH:5,5`-dithiobis(2-nitrobenzoic acid) (DTNB) oxidoreductase activities of all mutant enzymes were less than 3% of the activity of wild-type enzyme. The weak DTNB reductase activities in the mutant enzymes lacking Cys-337 or Cys-340 may occur through direct reduction of the mixed disulfide Cys-337-thiol or Cys-340-thiol and nitrothiobenzoate by FADH(2). However, the weak DTNB reductase activity in the mutant enzyme lacking both cysteines indicates that FADH(2) can reduce either DTNB or another disulfide directly, albeit inefficiently. These results suggest intramolecular dithiol-disulfide interchange reactions in the flavoprotein.


INTRODUCTION

A new group of facultatively anaerobic bacteria has been isolated from an alkaline compost and named Amphibacillus xylanus(1, 2) . Because A. xylanus metabolizing via either anaerobic or aerobic pathways produces similar amounts of ATP, the bacteria grow well and have the same growth rate and cell yield under both conditions in spite of lacking a respiratory system(2, 5) . We have proposed that NAD is regenerated by NADH oxidase from NADH produced from glycolysis and pyruvate oxidation in the aerobic pathway(5) .

A flavoprotein that is functional as NADH oxidase is purified from aerobically grown A. xylanus(3) . The flavoprotein is a homotetramer composed of a subunit (M(r) = 56,000) containing 1 mol of FAD and catalyzes the reduction of oxygen to hydrogen peroxide with beta-NADH as the preferred electron donor(3) . The enzyme is also observed to catalyze a thiol-disulfide interchange reaction, NADH:DTNB (^1)oxidoreductase(4) .

The NADH oxidase activity of the flavoprotein has a ping-pong type mechanism, and the K for NADH and the K for oxygen are 33.3 µM and 1.7 mM, respectively. The K value for oxygen is too high to catalyze the efficient reoxidization of NADH by oxygen in the cell of A. xylanus. In the presence of free FAD, however, the K value for oxygen becomes much lower so that the accurate determination of the value by the usual assay method is not possible. NADH oxidase activity is accelerated markedly in the presence of free FAD. The intracellular FAD concentration of A. xylanus is calculated to be about 13 µM, and the NADH oxidase activity is accelerated sufficiently with this FAD concentration. These results indicate that the flavoprotein acts as an effective NADH oxidase in the cell of A. xylanus(4) .

Dithionite and NADH titration of the flavoprotein require 3 eq of reductant/FAD for full reduction. During the addition of the first 4 electrons in the dithionite titration, the two bands of the oxidized flavin spectrum decrease gradually, and a new absorbance band at 585 nm forms. The long wavelength absorbance is ascribed to a neutral (blue) flavin semiquinone. The second phase of the reduction results in a further 2-electron reduction of the enzyme and spectral changes associated with full reduction of the flavin component. These results indicated that the flavoprotein has non-flavin redox center(s) in addition to the FAD(4) . Two disulfide bonds have been demonstrated in the flavoprotein. In order to explain the observed 6-electron reduction from titration, we proposed that the two disulfides act as the non-flavin redox centers(4) .

The amino acid sequence of A. xylanus flavoprotein is highly homologous with the alkyl hydroperoxide reductase F52a pro- tein component from Salmonella typhimurium(11) and the NADH dehydrogenase from an alkalophilic Bacillus sp. YN-1 (12) in all domains, showing identity of 51.2 and 72.5%, respectively(3) . Further, the short segment of the flavoprotein, which contained Cys-337 and Cys-340, shows a high degree of homology with Escherichia coli thioredoxin reductase(4) . The homologous region of thioredoxin reductase contains the redox-active disulfide, Cys-135 and Cys-138, which is active in catalysis(19, 20, 24) . Alkyl hydroperoxide reductase F52a protein is related to thioredoxin reductase, but there is an additional domain at the N terminus of alkyl hydroperoxide reductase F52a protein. Both enzymes contain FAD and a redox-active disulfide, in each monomer, in homologous positions. The N-terminal extension of alkyl hydroperoxide reductase F52a protein contains an additional disulfide that appears to be able to interchange with the redox-active disulfide(11, 13) . Therefore, we hypothesized that Cys-337 and Cys-340 of the flavoprotein might create a disulfide and be involved in the flow of electrons. To confirm our hypothesis, we have created three site-directed mutations of Cys-337 and Cys-340 of A. xylanus flavoprotein in an attempt to assign specific catalytic roles to these thiols. In this report, we describe the spectral and steady-state kinetic analysis of these altered enzymes.


EXPERIMENTAL PROCEDURES

Materials

DNA modification and restriction enzymes were the products of Takara Shuzo Co. (Kyoto, Japan). Butyl-Toyopearl and Toyopearl HW-60 were obtained from Tosoh (Tokyo, Japan). 5`-AMP Sepharose 4B and preswollen DE53 anion exchange resin were purchased from Pharmacia LKB Biotechnology Inc. and Whatman, respectively. FAD and beta-NADH were obtained from Sigma. All other chemicals were of reagent grade.

Bacterial Strains and Plasmids

Plasmid pNOH1850, which was designed for overexpression of the wild-type flavoprotein gene(4) , was used to construct derivatives that expressed the mutant flavoprotein. E. coli JM109 was used as host strains for the wild-type and mutant pNOH1850. The E. coli CJ236, BMH71-18 mutS, and MV1184 were used as directed in the site-directed mutagenesis system Mutan-K (Takara Shuzo Co.). E. coli cells were grown with shaking at 37 °C in 2 times YT or LB medium(8) .

Plasmid Constructions and Mutagenesis

Plasmid pNOH1850 was digested with SacI and ScaI, and the resulting 304-base pair fragment that includes Cys-337 and Cys-340 of the coding region of wild-type flavoprotein was purified from 1.0% agarose gel by using the GF/C disk (Whatman) as described by Chen and Thomas(6) . The 304-base pair fragment and SacI-SmaI cleaved pUC119 were mixed, ligated, and transformed into E. coli MV1184 to obtain single-stranded DNA using standard procedures (see Fig. 1)(32) .


Figure 1: Construction of mutant expression plasmid. Construction of the wild-type expression plasmid has been described(4) . The filled boxes represent the A. xylanus genes, and the open boxes represent the 1.9-kilobase HindIII fragments containing the sequence of the flavoprotein gene. The hatched boxes represent the tac promoters, and the cross-hatched boxes represent the Amp gene. Asterisks indicate the mutated codons. The position of the flavoprotein and the mutant flavoprotein gene giving rise to translated protein is indicated by the labeled arrow. kb, kilobase.



The mutagenic oligonucleotides used were 5`-TAGCATATTCTACACAC-3` (C337S), 5`-GTACACACTCCGATGCC-3` (C340S), and 5`TGTAGCATATTCTACACACTCCGATGCCCC-3` (C337S/C340S). Mutagenesis reactions were carried out as described by Kunkel (7) employing the site-directed mutagenesis system Mutan-K. Single-stranded pNOSS906 served as the template (Fig. 1). Double-stranded circular DNA resulting from the mutagenesis reactions was transformed into E. coli MV1184. The resulting DNA prepared from individual colonies was sequenced to verify the presence of the desired mutations. Dideoxy sequencing was performed using the BcaBEST dideoxy sequencing kit (Takara Shuzo Co.) according to the manufacturer's instructions with [alpha-P]dCTP (Amersham).

Replicative form DNA of phagemids carrying the mutations for C337S, C340S, and C337S/C340S was digested with BglII, and the resulting 216-base pair fragment was recovered from 1.5% agarose gel. The fragment was inserted into BglII-cleaved pNOH1850 to construct the mutant expression plasmids, and the resulting plasmids were transformed into E. coli JM109. Plasmid DNA was isolated by using standard alkaline lysis methods, and manipulation was performed according to standard methods(8) .

Purification of Mutant Enzymes

Three mutant enzymes were purified to homogeneity from E. coli JM109 harboring the respective mutant plasmids by the method described previously except for modification of the elution conditions of 5`-AMP Sepharose 4B chromatography(4) . The pool of active fractions after butyl-Toyopearl chromatography was concentrated and dialyzed against 2 times 4 liters of 50 mM potassium phosphate buffer, pH 6.0, containing 5 mM EDTA and 25 µM FAD. After centrifugation at 31,000 times g for 20 min, the supernatant was applied to a column of 5`-AMP Sepharose 4B (1.5 times 20 cm) equilibrated in 50 mM potassium phosphate buffer, pH 6.0, containing 5 mM EDTA. After the column was washed with 50 mM potassium phosphate buffer, pH 6.0, containing 5 mM EDTA and 25 µM FAD, the mutant enzyme was eluted with 50 mM potassium phosphate buffer, pH 6.0, containing 5 mM EDTA, 25 µM FAD, and 7.5 mM 5`-AMP. Purity of the mutant enzymes was judged by SDS-polyacrylamide gel (12.5%) electrophoresis (10) and spectroscopic analysis. Absorbance spectra were recorded with a Hitachi 557 double wavelength double beam spectrophotometer.

Extinction Coefficient Determination

The extinction coefficient of the protein bound FAD at 450 nm was determined for each of the flavoprotein species by resolving the FAD from the protein and quantitating the free FAD. Spectra of samples of the three proteins in 50 mM sodium phosphate buffer, pH 7.0, containing 0.5 mM EDTA were obtained, and the FAD was then released by the addition of 0.1% SDS (final concentration) at room temperature for 30 min.

Thiol and Disulfide Quantitations

Disulfides and thiols were quantitated spectrophotometrically as described previously(4) . Thiols were quantitated with DTNB under non-reducing and NADH-reducing conditions in 6 M guanidine hydrochloride at pH 8.0 by the method of Ellman(21) . Disulfide bonds were titrated with disodium 2-nitro-5-thiosulfobenzoate in excess sodium sulfite containing 2 M guanidine thiocyanate (final concentration) at pH 9.5 as described by Thannhauser et al.(9) .

Anaerobic Titrations

All anaerobic titrations with NADH and dithionite were performed as described previously (4) in double-side-armed anaerobic cuvettes similar to those of Williams et al.(14) . The anaerobic enzyme sample in 50 mM sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA was prepared by sequential evacuation and re-equilibration with oxygen-free argon. Oxygen-free argon was prepared by passing commercially obtained pure argon through a column of Oxyout (Osaka Sanso Co.). Dithionite solutions were prepared in the same buffer and standardized anaerobically against riboflavin before or after each set of enzyme titrations.

Steady-state Kinetics

Initial velocity studies of the NADH oxidase activity of the flavoprotein followed assay procedures described previously(4) . K(m) values for NADH and oxygen were determined from Lineweaver-Burk plots of kinetic data obtained at 25 °C by varying both NADH and oxygen concentration in 50 mM sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA.

NADH:DNTB Oxidoreductase Assay

Anaerobic enzyme samples (0.08-0.10 µM) in 50 mM sodium phosphate buffer, pH 7.5, containing 0.5 mM EDTA were prepared in anaerobic cuvettes equipped with two side arms by sequential evacuation and re-equilibration with oxygen-free argon. The reaction was started by mixing the NADH and DTNB at 25 °C, and the change of absorbance was recorded at 412 nm. The concentrations of NADH and DTNB for assay were 0.1 and 1 mM, respectively.

Thermal Stability

Wild-type and mutant enzymes were incubated for 10 min at several temperatures in the range of 4-70 °C. NADH oxidase activity was then measured at 37 °C in sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA. The concentration of NADH for the assay was 100 µM without additional FAD.

General Procedures

Other analytical methods followed protocols previously described(3, 4) .


RESULTS

Mutagenesis and Preparation of Mutant Enzyme

We have constructed three mutant expression plasmids using the mutagenic oligonucleotides and the expression system described previously(4) . All mutant enzymes were expressed at a level approximately equivalent to the wild-type enzyme. The purification scheme for all the enzymes is outlined under ``Experimental Procedures.'' SDS-polyacrylamide gel electrophoresis analysis of the purified mutant enzymes gave a single band having a molecular weight of 56,000. The absorbance ratio A/A is 5.2 for the wild-type enzyme and varies from 4.9 to 5.4 for C337S, C340S, and C337S/C340S mutants, corresponding to a maximal flavin loading of one FAD/subunit. No significant flavin loss was observed in these mutants even after dialysis, indicating that there is little or no disruption of the FAD binding domain caused by Cys-337 or Cys-340 mutations.

Spectral Analysis

The extinction coefficients in the 450 nm region of the protein-bound FAD of the three proteins were determined: C337S, = 13,100 M cm; C340S, = 12,800 M cm; and C337S/C340S, = 12,800 M cm. The absorbance spectra of oxidized wild-type and mutated enzymes were virtually identical (data not shown). The peaks for three mutant enzymes were located at 380 and 450 nm, the same as for wild-type enzyme. The shoulder in the 480 nm region, indicative of the hydrophobic environment of the flavin(15) , is unchanged in the mutant enzymes.

Thiol and Disulfide Quantitations

The mutations result in a change in the number of thiols and disulfide bridges in the protein. This change was confirmed by using DTNB and disodium 2-nitro-5-thiosulfobenzoate. For reference, there are, putatively, 2 disulfide bridges and 1 free cysteine residue in the wild-type enzyme. The number of thiols present in the enzymes was quantitated by reaction with DTNB under non-reducing conditions or in the presence of excess NADH in a final concentration of 6 M guanidine HCl at pH 8.0. 1 and 3.5 thiols of wild-type enzyme were reacted with DTNB in its oxidized and NADH-reduced form, respectively. This increase of thiols indicated complete reduction of the redox-active disulfide and partial reduction of a second disulfide in the wild-type enzyme, suggesting that there are two disulfides in the wild-type enzyme. By contrast, C337S, C340S, and C337S/C340S mutants exhibited 2.0 and 2.1, 1.9 and 2.0, and 1.0 and 1.1 thiols/subunit in their oxidized and NADH-reduced forms, respectively. The three enzymes were denatured, and the disulfides were quantitated by reacting with disodium 2-nitro-5-thiosulfobenzoate at pH 9.5 in the presence of sulfite. Only a single disulfide was found in each of the mutant enzymes. These results confirm that Cys-337 and Cys-340 contribute to a disulfide bridge and indicate that it is the redox-active disulfide in the wild-type enzyme.

Dithionite Titration of Mutant Enzymes

Wild-type flavoprotein can exist in three spectrally distinct redox states: fully oxidized, characterized by an absorbance maximum at 450 nm; 4-electron-reduced, possessing a contribution from neutral (blue) flavin semiquinone; and 6-electron-reduced, exhibiting a bleached spectrum due to reduction of FAD prosthetic groups(4) .

Titration of C337S, C340S, and C337S/C340S mutant enzymes with dithionite requires only 2 electron eq to reach the reduced flavin state (Fig. 2). These results show the absence of the redox-active disulfide and demonstrate the involvement of Cys-337 and Cys-340 in the redox-active disulfide. In the 1-electron-reduced state, the flavin absorbance of the oxidized C337S, C340S, and C337S/C340S mutant enzymes diminishes and a new band appears at around 580 nm. The new band, having a shoulder at 600 nm, is typical of the neutral (blue) flavin semiquinone(16, 18) , which is also observed in the 4-electron reduction of wild-type enzyme(4) . In the C340S and C337S/C340S mutant enzymes, the maximal formation of semiquinone is 45% of the total enzyme flavin. The absorbance values at 580 and 450 nm of 100% semiquinone were extrapolated from a plot of A versus A(18) , revealing that the extinction coefficients of flavin semiquinone are 4,800-4,900 M cm at 580 nm and 4,400-4,600 M cm at 450 nm in both mutant enzymes. These extinction coefficients are higher than that of wild-type enzyme (4,400 M cm and 4100 M cm, respectively). Because the absorption band of FAD at 450 nm did not change shape or peak position and because no absorption band around 750 nm was observed, this indicates there was no stable thiolate-flavin charge-transfer species(33) . In contrast, the maximal formation of semiquinone is 17% of the total enzyme in C337S mutant enzyme, and the extinction coefficient of this band is 3,700 M cm and 3,200 M cm at 580 and 450 nm, respectively. These values are lower than those of the wild-type enzyme. Further addition of dithionite reduces the remaining FAD and FADH to FADH(2).


Figure 2: Spectral titration of flavoprotein with dithionite. The NADH oxidase, 30.0-37.5 µM in 50 mM sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA was titrated at 20 °C with dithionite. Spectra were recorded after each addition when no further absorbance changes occurred. The spectra are of oxidized enzyme (top lines) and enzyme after reduction by dithionite. The insets show the absorbance at 450 nm versus added dithionite. A, C337S mutant enzyme; B, C340S mutant enzyme; C, C337S/C340S mutant enzyme.



NADH Titration of Mutant Enzymes

NADH titration of the three mutant enzymes required 1 eq of NADH/FAD (Fig. 3), but the titration had three apparent phases. Spectra typical of semiquinone formed in the first phase, much like the behavior upon dithionite titration. The accumulation of semiquinone with NADH titrations was the same as in titrations with dithionite for all of the mutant enzymes. In the second apparent phase, the formation of a new long wavelength absorbance band at 700-800 nm was observed in all mutant enzymes. The new band is the familiar FADH(2)-NAD charge transfer complex with its broad long wavelength absorbance extending beyond 800 nm(17) , which was also observed in NADH titrations of wild-type enzyme(4) . In the third apparent phase, the reduction is completed and excess NADH accumulates as shown by increased absorbance at 350 nm. The stability of the FADH(2)-NAD complex in the presence of excess NADH indicates very tight binding of NAD relative to NADH.


Figure 3: Spectral titration of flavoprotein with NADH. The NADH oxidase, 28.2-32.0 µM flavoprotein in 50 mM sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA was titrated at 20 °C with NADH. The spectra are of oxidized enzyme (top lines) and enzyme after reduction by NADH. The insets show the absorbance at 450 nm versus added NADH. A, C337S mutant enzyme; B, C340S mutant enzyme; C, C337S/C340S mutant enzyme.



Reduction of each of the mutant enzymes with NADH shows that only the FAD prosthetic group is involved. A comparison of these results with NADH titration of wild-type enzyme confirms that Cys-337 and Cys-340 constitute the active-site disulfide in direct communication with flavin. This redox pair in turn is presumably responsible for reduction of the remaining disulfide.

NADH Oxidase Activity of Mutant Enzyme

The catalytic activity of the three enzymes was examined by a steady-state analysis. The Lineweaver-Burk plots of initial velocities of all mutant enzymes gave parallel indications of a ping-pong type mechanism, as is the case with the wild-type enzyme. The replots of the y-intercepts versus [oxygen] were linear for all mutant enzymes (data not shown). The kinetic constants measured by this assay are given in Table 1. The values of K(m) for NADH and oxygen and the k values of these mutant enzymes were not different from those of the wild type.



NADH oxidase activity of wild-type enzyme was accelerated markedly in the presence of additional free FAD, but FMN and riboflavin did not affect the NADH oxidase activity(4) . The NADH oxidase activity of all mutant enzymes was accelerated markedly in the presence of free FAD (Fig. 4), as with the wild-type enzyme(4) . The optimum pH value for the activity of all mutant enzymes was not significantly altered relative to that of the wild type (data not shown). These results indicated that Cys-337 and Cys-340 of the flavoprotein are not involved in the NADH oxidase activity.


Figure 4: Effect of FAD on NADH oxidase activity. Assay conditions were 50 mM sodium phosphate buffer, pH 6.6, containing 0.5 mM EDTA and 100 µM NADH at 25 °C. Open circles, wild-type enzyme; filled circles, C337S; open squares, C340S; filled triangles, C337S/C340S.



NADH:DTNB Oxidoreductase Activity of Mutant Enzyme

The wild-type flavoprotein has been shown to catalyze electron transfer between NADH and DTNB(4) . The NADH:DTNB oxidoreductase activity in all mutant enzymes was measured at 25 °C in 50 mM sodium phosphate buffer (pH 7.5). All mutant enzymes were found to have less than 3% of the activity of wild-type enzyme in a NADH:DTNB oxidoreductase assay. Also, the DTNB reduction activity of all mutant enzymes was examined as a function of pH (4.5-8.0) using 100 mM citrate-Na(2)HPO(4) buffer containing 0.5 mM EDTA. All mutant enzymes had negligible NADH:DTNB oxidoreductase activity over the whole pH range. These results indicated that Cys-337 and Cys-340 are involved with the NADH:DTNB oxidoreductase activity in the wild-type enzyme.

Thermal Stability

In order to test if the replacements of Cys-337 and Cys-340 had induced any significant conformational change, all the mutant enzymes were incubated for 10 min at several different temperatures in the range of 4-70 °C. NADH oxidase activity was then measured at 37 °C. The K(m) for oxygen and the K(m) for NADH of the three mutant enzymes were not significantly altered relative to the wild-type enzyme. The mutant enzymes were all found to undergo cooperative thermal denaturation at the same temperature (approximately 60 °C) as wild-type enzyme. The lack of change in thermal stability strongly suggests that none of the mutations has led to a major conformational change in the enzymes and suggests that the proteins were not significantly different in threedimensional structure.


DISCUSSION

The A. xylanus flavoprotein that functions as NADH oxidase has unique functional properties that are the difference from known NADH oxidases(25, 26, 27, 28, 29, 30, 31) . Several enzymes referred to as NADH oxidase are known to be able to catalyze electron transfer from NADH to various electron acceptors such as methylene blue, cytochrome c, 2,6-dichloroindophenol, and potassium ferricyanide(26, 27, 30, 31) . However, none of these enzymes have been reported to be able to catalyze thiol-disulfide interchange reactions. A. xylanus flavoprotein was obtained to catalyze electron transfer between NADH and DTNB(4) . We have hypothesized that Cys-337 and Cys-340 of the flavoprotein are involved in the flow of electrons from NADH as the non-redox center(4) .

Pseudomonas aeruginosa mercuric reductase, which is a key component of an organomercurial detoxification system, has a redox-active disulfide (Cys-135,Cys-140) and a FAD(41) . Mercuric reductase partially reduced with NADPH is an EH(2) species whose spectral features have been attributed to a charge-transfer interaction between the thiolate of Cys-140 and FAD(41) . Further, the enzyme reduced by 0.8 eq of dithionite shows fluorescence intensity of the enzyme flavin diminished by 44% but only shows a 16% thiolate-FAD charge transfer, indicating that the second disulfide is in close proximity to the FAD(22) . Miller et al.(22) designated the Cys-558,Cys-559 disulfide as the auxiliary disulfide and suggested that reduction of the auxiliary disulfide by NADPH occurs via dithiol-disulfide interchange with the redox-active cysteine pair (Cys-135, Cys-140).

Reduction of the enzyme FAD by the strong reductant dithionite occurred during the total uptake of 6 electrons in the A. xylanus wild-type flavoprotein. Because the FAD can accept only 2 electrons, the 6-electron uptake indicates that other redox-active acceptors are present. The amino acid sequence suggests that these are disulfides (4) . Titration of C337S, C340S, and C337S/C340S mutant enzymes with NADH required only 2 electrons for full reduction. These results clearly indicate the absence of redox-active disulfide and demonstrate the involvement of Cys-337 and Cys-340 in the redox-active disulfide. This agrees with the results of thiol quantitation in that the number of thiols is the same in oxidized and reduced mutant enzymes. These results show that the redox-active disulfide, Cys-337 and Cys-340, is reduced directly via the 2-electron-reduced FAD coenzyme.

The NADH oxidase activity of wild-type enzyme is accelerated markedly in the presence of additional free FAD, and the K(m) value for oxygen decreases dramatically(4) . The K(m) values for oxygen and NADH and the k values in the three mutant enzymes were not significantly changed. The acceleration of NADH oxidase activity by excess FAD is also observed in all three mutant enzymes. The NADH oxidase activity of the mutant enzymes that lack the redox-active disulfide shows that the activity does not depend on the other disulfide. Thus, it is clear that neither disulfide bond in the wild-type enzyme is involved in the reduction of oxygen to hydrogen peroxide. This suggests that the reaction of 2-electron-reduced FAD coenzyme with oxygen may form a flavin-C-4a-hydroperoxide adduct(34) , followed by the elimination of hydrogen peroxide. Ahmed and Claiborne (35, 36) proposed that the peroxidatic reaction in Streptococcus faecalis NADH oxidase, which catalyzes the reduction of oxygen to water, in contrast, would involve the redox-active cysteinyl derivative.

The pyridine nucleotide-disulfide oxidoreductases form a family of homodimeric flavoproteins, having a redox-active disulfide and FAD in each monomer. Interaction of the 2electron-reduced enzymes involves sequential thiol-disulfide interchange reactions. Each nascent thiol of 2-electron-reduced enzyme has a distinct function; the interchange thiol reacts with the disulfide substrate, and the electron-transfer thiol reacts with the FAD(20) . Studies of two active site mutations of E. coli thioredoxin reductase, Ser-135,Cys-138 and Cys-135,Ser-138, have shown that Cys-138 interacts more closely with the FAD than does Cys-135(37, 38, 39) . This was confirmed by the study of x-ray crystal structure; Cys-138 is indeed close to the flavin, 3.0 Å from the C-4a position of the isoalloxazine ring, whereas Cys-135 is 4.4 Å from the C-5a position of the isoalloxazine ring(40) . It is suggested that Cys-135 might be the interchange thiol, because Cys-138 interacts with the FAD(20, 38, 39) .

The active site mutant of mercuric reductase from P. aeruginosa, Cys-135,Ser-140, shows 3% of the rate of DTNB reduction of the wild-type enzyme. Schultz et al. (42) suggested that the mixed disulfide between Cys-135-thiol and nitrothiobenzoate is reduced directly by FADH(2). Further, they suggest that the Ser-135,Cys-140 mutant enzyme lacks DTNB reductase activity due to steric hindrance in the formation of the mixed disulfide(42) .

All three mutant enzymes showed weak NADH:DTNB oxidoreductase activities, less than 3% of the rate of the wild-type enzyme. Thus, it is clear that Cys-337 and Cys-340 are involved with the NADH:DTNB oxidoreductase activity in the wild-type enzyme. These activities were the same in both mutant enzymes, suggesting that the thiol-disulfide interchange reactions differ from that of thioredoxin reductase, in spite of the high degree of conservation around the active site cysteines. The weak DTNB reduction in the C337S and C340S mutant enzymes may occur slowly through direct reduction by FADH(2) of the mixed disulfide between Cys-337-thiol or Cys-340-thiol and nitrothiobenzoate. Surprisingly, the C337S/C340S mutant enzyme, lacking both thiols, also showed the same 3% DTNB reductase activity. This result indicates that the other disulfide bond in the C337S/C340S mutant enzyme may also have thiol-disulfide interchange activity. We suggest that the thiol-disulfide interchange reaction in the wild-type flavoprotein may involve an intramolecular dithiol-disulfide interchange between the second disulfide and the Cys-337,Cys-340 thiols, reduced via FADH(2). This would differ from the known members of the pyridine nucleotide-disulfide oxidoreductase family of flavoprotein(20) , except for mercuric reductase(22) .


FOOTNOTES

*
This work was supported by grants from the Hayashida Foundation of the Tokyo University of Agriculture and from the Japanese Ministry of Education, Science, and Culture. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Food Science and Technology, Tokyo University of Agriculture, 196 Yasaka Abashiri-shi, Hokkaido, 099-24 Japan. Tel.: 81-0152-48-2116; Fax: 81-0152-48-2940.

(^1)
The abbreviations used are: DTNB, 5,5`-dithiobis(2-nitrobenzoic acid); C337S, flavoprotein mutant with serine at residue 337; C340S, flavoprotein mutant with serine at residue 340; C337S/C340S, flavoprotein mutant with serines at residues 337 and 340.


ACKNOWLEDGEMENTS

We thank Dr. Charles H. Williams, Jr. at the University of Michigan for several helpful suggestions, valuable advice, and critical reading of the manuscript. We thank Dr. Vincent Massey at the University of Michigan for helpful discussions and suggestions. We also thank Kengo Suzuki for assistance in analytical experiments.


REFERENCES

  1. Niimura, Y., Yanagida, F., Uchimura, T., Ohara, N., Suzuki, K., and Kozaki, M. (1987) Agric. Biol. Chem. 51, 2271-2275
  2. Niimura, Y., Yanagida, F., Suzuki, K., Komagata, K., and Kozaki, M. (1990) Int. J. Syst. Bacteriol. 40, 297-301
  3. Niimura, Y., Ohnishi, K., Yarita, Y., Hidaka, M., Masaki, H., Uchimura, T., Suzuki, H., and Kozaki, M. (1993) J. Bacteriol. 175, 7945-7950 [Abstract]
  4. Ohnishi, K., Niimura, Y., Yokoyama, K., Hidaka, M., Masaki, H., Uchimura, T., Suzuki, H., Uozumi, T., Kozaki, M., Komagata, K., and Nishino, T. (1994) J. Biol. Chem. 269, 31418-31423 [Abstract/Free Full Text]
  5. Niimura, Y., Koh, E., Uchimura, T., Ohara, N., and Kozaki, M. (1989) FEMS Microbiol. Lett. 61, 79-84
  6. Chen, W. C., and Thomas, C. A., Jr. (1980) Anal. Biochem. 101, 339-341 [Medline] [Order article via Infotrieve]
  7. Kunkel, K. T. (1985) Proc. Natl. Acad. Sci. U. S. A. 82, 488-492 [Abstract]
  8. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , 2nd Ed., Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  9. Thannhauser, T. W., Konishi, Y., and Scheraga, H. A. (1984) Anal. Biochem. 138, 181-188 [Medline] [Order article via Infotrieve]
  10. Laemmli, U. K. (1970) Nature 227, 680-685 [Medline] [Order article via Infotrieve]
  11. Tartaglia, L. A., Storz, G., Brodsky, M. H., Lai, A., and Ames, B. N. (1990) J. Biol. Chem. 265, 10535-10540 [Abstract/Free Full Text]
  12. Xuemin, X., Koyama, N., Cui, M., Yamagishi, A., Nosoh, Y., and Oshima, T. (1991) J. Biochem. (Tokyo) 109, 678-683 [Abstract]
  13. Jacobson, F. S., Morgan, R. W., Christman, M. F., and Ames, B. N. (1989) J. Biol. Chem. 264, 1488-1496 [Abstract/Free Full Text]
  14. Williams, C. H., Jr., Arscott, L. D., Matthews, R. G., Thorpe, C., and Wilkinson, K. D. (1979) Methods Enzymol. 62, 185-198 [Medline] [Order article via Infotrieve]
  15. Harbury, H. A., LaNoue, K. F., Loach, P. A., and Amick, R. M. (1959) Proc. Natl. Acad. Sci. U. S. A. 45, 1708-1711
  16. Massey, V., and Palmer G. (1966) Biochemistry 5, 3181-3189 [Medline] [Order article via Infotrieve]
  17. Massey, V., and Palmer, G. (1962) J. Biol. Chem. 237, 2347-2358 [Free Full Text]
  18. Zanetti, G., Williams, C. H., Jr., and Massey, V. (1968) J. Biol. Chem. 243, 4013-4019 [Abstract/Free Full Text]
  19. Russel, M., and Model, P. (1988) J. Biol. Chem. 263, 9015-9019 [Abstract/Free Full Text]
  20. Williams, C. H., Jr. (1992) in Chemistry and Biochemistry of Flavoenzymes (M ü ller, F., ed) Vol. 3, pp. 121-209, CRC Press, Boca Raton, FL
  21. Ellman, G. L. (1959) Arch. Biochem. Biophys. 82, 70-77 [Medline] [Order article via Infotrieve]
  22. Miller, S. M., Moore, M. J., Massey, V., Williams, C. H., Jr., Distefano, M. D., Ballou, D. P., and Walsh, C. T. (1989) Biochemistry 28, 1194-1205 [Medline] [Order article via Infotrieve]
  23. Deleted in proof
  24. Ronchi, S., and Williams, C. H., Jr. (1972) J. Biol. Chem. 247, 2083-2086 [Abstract/Free Full Text]
  25. Schmidt, H. L., Stocklein, W., Danzer, J., Kirch, P., and Limbach, B. (1986) Eur. J. Biochem. 156, 149-155 [Abstract]
  26. Park, H. J., Reiser, C. O. A., Kondoruweit, S., Erdomann, H., Schmidt, R., and Sprinzl, M. (1992) Eur. J. Biochem. 205, 881-885 [Abstract]
  27. Koike, K., Kobayashi, S., Ito, S., and Saitoh, M. (1985) J. Biochem. (Tokyo) 97, 1279-1288 [Abstract]
  28. Cocco, D., Rinaldi, A., Savini, I., Cooper, J. M., and Bannister, V. (1981) Eur. J. Biochem. 174, 267-271 [Abstract]
  29. Reinards, R., Kubicki, J., and Ohlenbusch, H. (1981) Eur. J. Biochem. 120, 329-337 [Abstract]
  30. Saeki, Y., Nozaki, M., and Matsumoto, K. (1985) J. Biochem. (Tokyo) 98, 1433-1440 [Abstract]
  31. Higuchi, M., Shimada, M., Yamamoto, Y., Hayashi, T., Koga, T., and Kamio, Y. (1993) J. Gen. Microbiol. 139, 2343-2351 [Medline] [Order article via Infotrieve]
  32. Vieira, J., and Messing, J. (1987) Methods Enzymol. 153, 3-11 [Medline] [Order article via Infotrieve]
  33. Williams, C. H., Jr. (1965) J. Biol. Chem. 240, 4793-4800 [Free Full Text]
  34. Entsch, B., Ballou, D. P., and Massey, V. (1976) J. Biol. Chem. 251, 2550-2563 [Abstract]
  35. Ahmed, S. A., and Claiborne, A. (1989) J. Biol. Chem. 264, 19856-19863 [Abstract/Free Full Text]
  36. Ahmed, S. A., and Claiborne, A. (1989) J. Biol. Chem. 264, 19864-19870 [Abstract/Free Full Text]
  37. Prongay, A. J., Engelke, D. R., and Williams, C. H., Jr. (1989) J. Biol. Chem. 264, 2656-2664 [Abstract/Free Full Text]
  38. Prongay, A. J., and Williams, C. H., Jr. (1990) J. Biol. Chem. 265, 18968-18975 [Abstract/Free Full Text]
  39. Prongay, A. J., and Williams, C. H., Jr. (1992) J. Biol. Chem. 267, 25181-25188 [Abstract/Free Full Text]
  40. Waksman, G., Krishna, T. S. R., Williams, C. H., Jr., and Kuriyan, J. (1994) J. Mol. Biol. 236, 800-816 [CrossRef][Medline] [Order article via Infotrieve]
  41. Fox, B., and Walsh, C. T. (1982) J. Biol. Chem. 257, 2498-2503 [Abstract/Free Full Text]
  42. Schultz, P. G., Au, K. G., and Walsh, C. T. (1985) Biochemistry 24, 6840-6848 [Medline] [Order article via Infotrieve]

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