Arginine 91 Is Not Essential for Flavin Incorporation in Hepatic Cytochrome b5 Reductase

Archives of Biochemistry and Biophysics Vol. 389, No. 2, May 15, pp. 223–233, 2001 doi:10.1006/abbi.2001.2340, available online at http://www.idealibrary.com on

Arginine 91 Is Not Essential for Flavin Incorporation in Hepatic Cytochrome b 5 Reductase Christopher C. Marohnic and Michael J. Barber* ,1 Department of Biochemistry and Molecular Biology, University of South Florida, College of Medicine, and *H. Lee Moffitt Cancer Center and Research Institute, Tampa, Florida 33612

Received November 30, 2000, and in revised form February 26, 2001; published online April 24, 2001

Cytochrome b 5 reductase (cb5r) catalyzes the transfer of reducing equivalents from NADH to cytochrome b 5. Utilizing an efficient heterologous expression system that produces a histidine-tagged form of the hydrophilic, diaphorase domain of the enzyme, site-directed mutagenesis has been used to generate cb5r mutants with substitutions at position 91 in the primary sequence. Arginine 91 is an important residue in binding the FAD prosthetic group and part of a conserved “RxY STxx NS” sequence motif that is omnipresent in the “ferredoxin:NADP ⴙ reductase” family of flavoproteins. Arginine 91 was replaced with K, L, A, P, D, Q, and H residues, respectively, and all the mutant proteins purified to homogeneity. Individual mutants were expressed with variable efficiency and all exhibited molecular masses of approximately 32 kDa. With the exception of R91H, all the mutants retained visible absorption spectra typical of a flavoprotein, the former being produced as an apoprotein. Visible absorption spectra of R91A, L, and P were red shifted with maxima at 458 nm, while CD spectra indicated an altered FAD environment for all the mutants except R91K. Fluorescence spectra showed a reduced degree of intrinsic flavin fluorescence quenching for the R91K, A, and P, mutants, while thermal stability studies suggested all the mutants, except R91K, were somewhat less stable than the wild-type domain. Initialrate kinetic measurements demonstrated that the mutants exhibited decreased NADH:ferricyanide reductase activity with the R91P mutant retaining the lowest activity, corresponding to a k cat of 283 s ⴚ1 and a NADH Km of 105 ␮M, when compared to the wild-type doNADH main (k cat ⴝ 800 s ⴚ1 K m ⴝ 6 ␮M). These results demonstrate that R91 is not essential for FAD binding in cb5r; however, mutation of R91 perturbs the flavin

1 To whom correspondence should be addressed. Fax: (813) 9747357. E-mail: [email protected].

0003-9861/01 $35.00 Copyright © 2001 by Academic Press All rights of reproduction in any form reserved.

environment and alters both diaphorase substrate recognition and utilization. © 2001 Academic Press

Hepatic cytochrome b 5 reductase (cb5r, 2 EC 1.6.2.2) catalyzes the transfer of two reducing equivalents from the physiological electron donor, NADH, to two molecules of cytochrome b 5 (1, 2). The generation of reduced cytochrome b 5 represents a critical step in a variety of oxidation–reduction reactions that occur on the cytoplasmic surface of the endoplasmic reticulum. Thus, cytochrome b 5 reductase participates in a number of important processes, including the elongation and desaturation of fatty acids (3, 4), an intermediate step in cholesterol biosynthesis catalyzed by delta 7-sterol 5-desaturase (5) and the transfer of reducing equivalents to cytochrome P450 for subsequent use in the hydroxylation of steroid hormones and xenobiotic metabolism (6). In addition to the microsomal form, a soluble derivative of cb5r, identified as the product of an alternative transcript in reticulocytes (7), has been demonstrated in circulating erythrocytes, where it participates in the reduction of methemoglobin to hemoglobin (8). Microsomal cytochrome b 5 reductase has been isolated from a variety of mammalian sources including human (9), bovine (10), porcine (11), and rat (12) liver 2 Abbreviations used: cb5r, cytochrome b 5 reductase; FNR, ferredoxin:NADP ⫹ reductase; PDR, phthalate dioxygen reductase; NADH:FR, NADH:ferricyanide reductase; NADH:BR, NADH:cytochrome b 5 reductase; SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis; PCR, polymerase chain reaction; IPTG, isopropyl ␤-D-thioglucopyranoside; FPLC, fast protein liquid chromatography; TB, terrific broth; MALDI-TOF, matrix-assisted laser desorption ionization-time of flight; m/z, mass/charge ratio; ␮, ionic strength; Mops, 3-(N-morpholino)propanesulfonic acid; CD, circular dichroism; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; DMSO, dimethylsulfoxide.

223

224

MAROHNIC AND BARBER

and in all cases the enzyme has been demonstrated to be a monomeric, amphipathic protein (molecular mass approximately 35 kDa) composed of two discrete domains. A small hydrophobic N-terminal membraneanchoring domain (residues 1–28, M r approximately 3.1 kDa) which serves to orient the catalytic site at the membrane-aqueous interface and to facilitate rapid intermolecular electron transfer and a large hydrophilic catalytic or “diaphorase” domain (residues 29 –300, M r approximately 31.5 kDa) which contains a single noncovalent FAD prosthetic group. Separation of the two domains by limited proteolysis has facilitated isolation of a truncated, soluble form of the reductase which retained NADH:cytochrome b 5 reductase activity (13). In addition to the physiological NADH:cytochrome b 5 reductase activity, the enzyme can also utilize a variety of artificial electron acceptors, such as ferricyanide, MTT, and molecular oxygen, though with greatly varying efficiencies. Kinetic characterizations using either cytochrome b 5 or ferricyanide have demonstrated that the rate-limiting step with either acceptor involves hydride transfer from NADH to the enzyme-bound FAD followed by more rapid electron transfer to the oxidizing substrate (14, 15). Primary sequence alignments have identified cytochrome b 5 reductase as a member of a class of prokaryotic and eukaryotic flavoprotein oxidoreductases collectively referred to as the “FNR family.” Typified by the simple flavoprotein, ferredoxin:NADP ⫹ reductase (EC 1.18.1.2) (16), the FNR family has expanded to include portions of more complex flavoproteins, such as the FAD-containing domains of cytochrome P450 reductase (EC 1.6.2.4) (17), nitric oxide synthetase (EC 1.14.13.39) (18), methionine synthase reductase (EC 2.1.1.135) (19), assimilatory NADH:nitrate reductase (EC 1.6.6.1) (20) and dissimilatory sulfite reductase (EC 1.8.1.2) (21), the flavin-containing subunit of methane monooxygenase (EC 1.14.13.25) (22), and the FMN-containing protein phthalate dioxygen reductase (EC 1.14.12.7) (23). These enzymes comprise a diverse array of flavoproteins that participate in a variety of critical metabolic pathways that include steroid biosynthesis, xenobiotic detoxification, fatty acid desaturation and inorganic nitrate and sulfite reduction or exhibit roles in signal transduction and antioxidant protection. Multiple sequence alignments of FNR family members have indicated the presence of several regions of highly conserved primary structure including a FADdomain carboxy-terminal sequence “motif” corresponding to the sequence “RxY STxx NS”. Comparative studies of the FNR and PDR X-ray structures have indicated that this motif is involved in flavin binding within these proteins, including interactions with the isoalloxazine ring and the FMN- and AMP-linked phosphate groups (24).

For rat cb5r, the amino acid residues comprising the “RxY STxx NS” motif correspond to the amino acids R91 3 to S97 within the mature protein and encompass the sequence “RPYTPVS.” Preliminary studies of the functional roles of four of these amino acids in cb5r, corresponding to R91, Y93, T94, and S97, using alaninescanning site-directed mutagenesis has indicated that substitutions of residues R91 and Y93 resulted in significantly diminished catalytic activity and small perturbations of the flavin absorbance spectrum, which suggested changes in FAD binding, while, in contrast, alanine substitutions of T94 and S97 had no effect on flavin binding and only modestly decreased cb5r catalytic activity (25). The X-ray structures of both porcine (26) and rat (27) cb5r indicate that R91, the initial residue of the motif, is situated toward the end of a ␤ strand (␤4) within the flavin-binding subdomain and opposes the si-face of the isoalloxazine ring of the FAD prosthetic group. The side-chain carbon atoms are oriented parallel to the FMN ribose and the guanidinium group is within hydrogen-bonding distance of both the FMN and AMP phosphate groups. We have generated a series of R91 mutants to examine the effects of specific amino acid substitutions at this position on both FAD prosthetic group binding and the interaction of the enzyme with the physiological substrate, NADH. MATERIALS AND METHODS Materials. Oligonucleotide primers were obtained from Integrated DNA Technologies (Coralville, IA). Native Pfu and Pfu Turbo polymerases as well as Epicurian coli BL21(DE3)-RIL cells were obtained from Stratagene (La Jolla, CA). Restriction enzymes (NdeI, BamHI, DpnI, HinFI, and BsmFI) were purchased from New England Biolabs (Beverly, MA). The pET23b vector was purchased from Novagen (Madison, WI) and T4 DNA ligase was purchased from Promega (Madison, WI). Triton X-100 and Hot Start Micro 50 PCR tubes were obtained from Molecular-Bio Products Inc. (San Diego, CA). Tryptone and yeast extract were obtained from EM Science (Gibbstown, NJ) and “Ultrol” grade Mops buffer was purchased from Calbiochem (La Jolla, CA). Sinapinic acid (3,5-dimethoxy-4-hydroxy cinnamic acid) was purchased from Aldrich Chemical Co. (Milwaukee, WI) and IPTG was obtained from RPI (Mt. Prospect, IL). Reagents for bacterial culture, protein purification, and chemical assays including NADH, NAD ⫹, NADPH, 5⬘-ADP-agarose, cytochrome c (horse heart, type VI), myoglobin (sperm whale), bovine serum albumin, riboflavin, FAD, K 3Fe(CN) 6, and trifluoroacetic acid were obtained from Sigma Chemical Co. (St. Louis, MO). Ni-NTA agarose and kits for plasmid preparation and agarose gel extraction were purchased from Qiagen Inc. (Valencia, CA). Nucleotide sequencing was performed by the Molecular Biology Core Facility at the H. Lee Moffitt Cancer Center and Research Institute. Construction of the pH4CB5R expression plasmid. The expression plasmid pH4CB5R was constructed using oligonucleotide prim-

3 Amino acid residues are numbered with respect to their position in the sequence of the full-length, membrane-binding form of cytochrome b 5 reductase.

225

CYTOCHROME b 5 REDUCTASE MUTAGENESIS TABLE I

Oligonucleotide Primers Used in the Construction of the Various H 4cb 5r R91 Mutants Protein R91K R91A R91L R91P R91D R91Q R91H WT

Primer (5⬘–3⬘) 5⬘-GGCAACTTGGTGATTCGTCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTGATTGCGCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTCTTCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTCCTCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTGATCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTCAGCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTCATCCCTACACCCCTGTG-3⬘ 5⬘-GGCAACTTGGTCATTCGTCCCTACACCCCTGTG-3⬘ N- G N L V I R P Y T P V -C

ers designed to specifically amplify the 810-bp region of the construct PGBK3 (27) corresponding to the coding sequence for the C-terminal 269-amino-acid residues of the rat cb5r sequence that comprise the soluble catalytic domain: Primer 1 (amino-terminus)—(5⬘–3⬘) CAT ATG CAC CAT CAT CAC ATG ATC ACC CTC GAG; Primer 2 (carboxy terminus)—(5⬘–3⬘) CGC GGA TCC CCG TCA GAA GGT GAA GCA TCG. Primer 1 also contained the sequence encoding an amino-terminal four-histidine affinity tag and a unique NdeI site (CAT ATG) for cloning the fragment into the pET23b expression vector. Primer 2 contained a stop codon and a unique BamHI (G GATCC) site for direct cloning of the fragment into the pET23b expression vector. PCR amplifications were performed using native Pfu polymerase (2.5 units) in the presence of native Pfu buffer containing 100 ng of template pGBK3 vector DNA, 1 ␮g of each synthetic primer, 5% DMSO, 0.1% Triton X-100, and 200 ␮M dNTPs. Initial melting was for 10 min at 94°C. Cycling parameters (32 cycles) were as follows: 1 min at 94°C, 1 min at 50°C, 1 min at 72°C. The amplified product was isolated by agarose gel electrophoresis and purified using the Qiagen gel extraction kit. Following digestion with NdeI and BamHI, the purified PCR fragment was cloned directly into the NdeI–BamHIdigested pET23b expression vector by ligation with T4 DNA ligase to create the construct pH4CB5R. The fidelity of the construct was verified by nucleotide sequencing in both directions. Site-directed mutagenesis. The pH4CB5R expression construct was specifically mutagenized using a modification of the Stratagene QuikChange (La Jolla, CA) protocol. Complimentary oligonucleotide primers (30 –36 mers) containing the desired codon change and a silent mutation (either inserting or deleting a restriction enzyme recognition sequence) were synthesized as shown in Table I. Vector PCR was performed using Pfu Turbo polymerase (1.25 units) in the presence of cloned Pfu buffer containing 10 ng pH4CB5R vector DNA, 125 ng of each synthetic primer, 5% DMSO, 0.1% Triton X-100, and 200 ␮M dNTPs with cycling parameters (18 cycles) of 1 min at 94°C, 1 min at 55°C, 10 min at 68°C. DpnI digestion of the reaction mixture following PCR was utilized to cleave only methylated template DNA and the PCR products were subsequently purified by agarose gel electrophoresis, melted for 5 min at 80°C, allowed to reanneal by gradient cooling to 4°C over 15 min, and used for direct transformation of competent Escherichia coli DH5␣ cells. Colonies were screened for the specific mutation by restriction enzyme digestion and visualization of the products by acrylamide gel electrophoresis. The fidelity of the mutant constructs were verified by nucleotide sequencing in both the forward and reverse directions. Positive constructs were then used to transform competent E. coli BL21(DE3)RIL cells. Cell culture and protein purification. E. coli BL21(DE3)-RIL cells harboring the pH4CB5R or mutant constructs were grown aerobi-

(⫹HinFI) (⫺BsmFI) (⫺BsmFI) (⫺BsmFI) (⫺BsmFI) (⫺BsmFI) (⫺BsmFI)

cally in TB media supplemented with riboflavin (100 ␮M) overnight at 37°C. Recombinant protein expression was induced by addition of IPTG (0.4 mM) and the cells cultured for an additional 6 h at 26°C. The cells were pelleted by centrifugation (400g, 10 min), resuspended in lysis buffer, comprising 50 mM Tris–HCl, containing 300 mm NaCl and 10 mM imidazole, pH 8.0, and disrupted by sonication in the presence of PMSF (1.2 mg/ml). Lysates were clarified by centrifugation (30,000g, 30 min) at 4°C and then incubated by gently rocking with Ni-NTA agarose (1 ml matrix/10 ml lysate) for up to 30 min at 4°C. The His-tagged-cb 5r-Ni-NTA matrix suspension was collected by centrifugation (180g, 5 min), washed twice with 25 mM phosphate buffer, containing 300 mM NaCl and 10 mM imidazole, pH 8.0, and transferred to a chromatography column (2.5 ⫻ 10 cm). Bound proteins were eluted with 25 mM phosphate buffer, containing 300 mM NaCl and 100 mM imidazole, pH 8.0. Yellow fractions containing cb5r were pooled, assayed for NADH:FR activity and concentrated by pressure filtration (Amicon YM5 membrane). Final purification was achieved by size-exclusion FPLC using a Superdex 200 column (1 ⫻ 30 cm) in 10 mM Mops buffer, containing 0.1 mM EDTA, pH 7.0. Fractions with NADH:FR activity containing cb5r were pooled, concentrated, beaded, and stored under liquid N 2 until required. Non-histidine-tagged cb5r was obtained as described by Barber and Quinn (28). The recombinant soluble, heme domain of rat cytochrome b 5 was produced as described by Beck-von Bodman et al. (29). Wild-type cb 5r concentrations were estimated using A 461 ⫽ 10.6 cm ⫺1 mM ⫺1. SDS–polyacrylamide gel electrophoresis was performed as described by Laemmli (30). MALDI-TOF mass spectrometry was performed as described by Barber and Quinn (28). Spectroscopy. UV/visible spectra were obtained using a Hewlett Packard (Agilent Technologies, Palo Alto, CA) 8453 diode-array spectrophotometer. UV and visible CD spectra were obtained using a JASCO (Easton, MD) J710 spectropolarimeter calibrated for both signal intensity and wavelength maxima using an aqueous solution of d-10-camphosulfonic acid (31). UV CD spectra were obtained in 10 mM phosphate buffer, containing 0.1 mM EDTA, pH 7.0, using a cylindrical quartz cell of 0.1 cm path length (300 ␮l total volume), while visible CD spectra were obtained in 10 mM Mops buffer, containing 0.1 mM EDTA, pH 7.0, using a cell of 1 cm path length (90 ␮l total volume). All spectra were corrected for the appropriate buffer contributions and are expressed in terms of molar ellipticities (M ⫺1 cm ⫺1). Fluorescence spectra were obtained using a Shimadzu Scientific Inst. Inc. RF-5301PC spectrofluorophotometer. Excitation and emission spectra were obtained using a slit width of 3 nm and emission and excitation wavelengths of 520 and 450 nm, respectively. Following acquisition of the wild-type and R91 mutant spectra, the enzymes samples were heated to 100°C for 30 min, centrifuged to remove protein, and the corresponding spectra for the liberated FAD

226

MAROHNIC AND BARBER

FIG. 1. SDS–polyacrylamide gel electrophoresis cb5r and the R91 mutants. Wild-type cb5r and the various R91 mutants were isolated as described under Materials and Methods and analyzed on a 15% polyacrylamide gel. Lane A, non-his-tagged cb5r (2 ␮g); lane B, H 4-tagged cb5r (2 ␮g); lane C, R91K (2 ␮g); lane D, R91A (2 ␮g); lane E, R91L (2 ␮g); lane F, R91P (2 ␮g); lane G, R91D (2 ␮g); lane H, R91Q (2 ␮g); lane I, R91H (2 ␮g). Lane S corresponds to protein molecular weight standards with the indicated molecular masses.

recorded. All spectra were corrected for the appropriate buffer contribution. Enzyme activities. NADH:FR, NADPH:FR, and NADH:BR activities were determined at 25°C under conditions of constant ionic strength and pH as previously described (32) in 116 mM Mops buffer (␮ ⫽ 0.05), containing 0.1 mM EDTA, pH 7.0. NAD(P)H:FR activities were routinely determined as the decrease in absorbance at 340 nm in the presence of NAD(P)H (100 ␮M) and ferricyanide (200 ␮M). NADH:BR activity was determined as the increase in absorbance at 423 nm in the presence of NADH (100 mM) and soluble, recombinant rat cytochrome b 5 (50 ␮M). Activities are expressed as initial rates for the oxidation of NAD(P)H (␮mol of NAD(P)H consumed/min/nmol FAD). Initial rate data were analyzed using the software “ENZFIT” (Elsevier Biosoft, Ferguson, MO) to yield apparent V max and K m values. Thermal stability measurements. Thermal stabilities of the wildtype and R91 mutant proteins were determined by monitoring both the release of the FAD prosthetic group, indicated by the increase in intrinsic flavin fluorescence and the loss of NADH:FR activity as previously described (33). Chemical modification of arginine residues. Chemical modification of arginine residues in both cb5r and the R91A mutant was performed at 23°C using either phenylglyoxal in 100 mM bicarbonate buffer, containing 0.1 mM EDTA, pH 8.1, or 2,3-butandione in 50 mM borate buffer, containing 0.1 mM EDTA, pH 8.0. Stock solutions of phenylglyoxal and 2,3-butanedione were freshly prepared in bicarbonate and borate buffer, respectively. Control samples were incubated under identical conditions in the absence of phenylglyoxal or 2,3-butanedione. NADH:FR activities were assayed in triplicate at the indicated time points and the activities expressed relative to a control sample maintained at 0°C. Phenylglyoxal-modified cb5r was prepared by incubating the enzyme (500 ␮M FAD) with phenylglyoxal (5 mM) for 5 min at 23°C in 100 mM bicarbonate buffer, containing 0.1 mM EDTA, pH 8.1, following which L-arginine was added to a final concentration of 200 mM and the modified enzyme isolated by FPLC using a Superdex 200 column in 10 mM Mops buffer, containing 0.1 mM EDTA, pH 7.0.

RESULTS

Site-directed mutagenesis was utilized to generate a series of substituted forms of histidine-tagged cb5r corresponding to R91K, R91A, R91L, R91P, R91D, R91Q, and R91H, respectively, where R91 corresponds to the initial residue in the “RxY STxx NS” motif (24) involved in

binding the FAD prosthetic group in cb5r. Initial attempts to generate the R91A mutant in the non-histidine-tagged form (28) proved unsatisfactory due to difficulties in purifying the mutant protein to homogeneity using the traditional isolation protocol involving ADP-agarose affinity chromatography (28), suggesting the mutant had a decreased affinity for the competitive inhibitor, 5⬘-ADP. However, using the his-tagged expression vector, all the mutants were successfully isolated using the standard NTA-agarose affinity chromatography and size exclusion FPLC protocol. Expression of the R91 mutants varied in efficiency potentially reflecting alterations in their intrinsic stability. R91L was expressed with the highest efficiency, comparable to levels obtained for the wild-type domain, while the R91H mutant was expressed at the lowest level and was observed to be the least stable. All the mutants were purified to homogeneity and analyzed by SDS– PAGE, as shown in Fig. 1, and in the low-mass range by MALDI-TOF mass spectrometry to verify FAD incorporation. These results indicated that all the mutants exhibited comparable molecular weights, of approximately 31 kDa, and all contained, with the exception of R91H, FAD as the sole prosthetic group. Representative UV/visible absorption spectra obtained for the oxidized forms of the wild-type cb5r and selected R91 mutants are shown in Fig. 2. The spectrum obtained for the R91K mutant was comparable to that obtained for the wild-type domain with absorption maxima at 276, 386, and 461 nm, respectively, together with a pronounced shoulder at 485 nm. However, for the remaining mutants, the visible absorption maxima were red shifted to approximately 458 – 459 nm with the greatest perturbations observed for the R91A,

FIG. 2. UV/visible absorption spectra of cb5r and R91 mutants. UV/visible absorption spectra were recorded for oxidized samples of the H 4-tagged wild type and the various cb5r R91 mutants at equivalent flavin concentrations (14 ␮M FAD) in 10 mM Mops buffer, containing 0.1 mM EDTA, pH 7.0. Individual spectra correspond to wild-type domain (—), R91K (— —), R91A (– –), R91L (- - -), R91P (. . .), and R91H (- 䡠 䡠 - 䡠 䡠 -).

CYTOCHROME b 5 REDUCTASE MUTAGENESIS

R91L, and R91P mutants. While several of the amino acid substitutions had small effects on the positions of the visible absorption maxima, all the FAD-containing mutants exhibited extinction coefficients at 458 – 461 nm comparable to the wild-type domain. In addition, the consistency of the A 276 /A 460 ratios obtained for the wild-type domain and the R91K, A, L, Q, D, and P mutants indicated no changes in the FAD stoichiometry for the mutant proteins. In contrast to the other mutants, the R91H variant did not retain a spectrum typical of a flavoprotein. While an absorption peak was apparent in the spectrum in the UV region at 276 nm, no absorption peaks were observed in the visible spectrum, indicating expression of the corresponding apoprotein. Various attempts to reconstitute the apoprotein with exogenous FAD were unsuccessful. To compare the folding of the wild type and R91 mutant proteins and to accurately assess the effects of the various amino acid substitutions on the spectroscopic properties of the FAD prosthetic group, CD spectra were obtained for the various oxidized proteins. Both far-UV and visible CD spectra are shown for the wild-type H 4cb 5r and the R91 mutants in Fig. 3. In the far-UV region, all the mutant proteins exhibited CD spectra comparable to that of the wild-type domain with a positive CD maximum in the range 196 –198 nm and a negative CD maximum in the range 218 –220 nm, indicative of no substantial perturbations of the overall secondary structure folding patterns for any of the flavin-containing mutants. In contrast, visible CD spectra obtained for the oxidized forms of the various flavin-containing R91 mutants, with the exception of the spectrum obtained for the R91K variant, showed considerable deviation from the spectrum obtained for the wild-type domain. The spectrum of the R91K mutant exhibited both positive and negative CD maxima at 310 and 390 nm and 460 and 485 nm, respectively, which were identical to the values obtained for the wild-type domain and of comparable spectral intensity. For the remaining R91 mutants, while the wavelengths corresponding to the various CD maxima were not significantly altered from the wild-type parameters, their polarities were reversed and their corresponding spectral intensities were substantially decreased, in the order R91D ⬎ R91L ⬎ R91A ⬎ R91P ⬎ R91Q. These perturbations were most significant for the R91Q mutant, which showed a spectrum more similar to free FAD than to the wild-type domain. Changes in the visible spectroscopic properties of the R91 mutants were also reflected in both the corresponding fluorescence excitation and emission spectra. Fluorescence excitation and emission spectra obtained for the wild-type domain and selected R91 mutants are shown in Fig. 4. Excitation spectra obtained for the wild type, R91K, R91D, and R91P mutants exhibited

227

FIG. 3. CD spectra of oxidized cb5r and R91 mutants. (Top) UV CD spectra were recorded using enzyme samples (7 ␮M FAD) in 10 mM phosphate buffer, containing 0.1 mM EDTA, pH 7.0, and correspond to H 4-tagged wild type domain (—); R91K (— —), R91A (– –), R91L (- - -), R91Q (- 䡠 䡠 - 䡠 䡠 -), R91D (- 䡠 - 䡠 - 䡠 ), and R91P (. . .). (Bottom) Visible CD spectra were recorded using enzyme samples (50 ␮M FAD) in 10 mM Mops buffer, containing 0.1 mM EDTA, pH 7.0, and correspond to H 4-tagged wild-type domain (—); R91K (— —), R91A (– –), R91L (- - -), R91Q (- 䡠 䡠 -), R91D (- 䡠 - 䡠 -), and R91P (. . .). The visible CD spectrum of a sample of free FAD (-. . .-. . .-) is shown for comparison.

maxima at 386 and 460 nm while the corresponding emission spectra yielded maxima at 515 nm. While the mutants showed only small shifts in the excitation and emission maxima, the degree of quenching of the intrinsic flavin fluorescence was reduced to 93, 88, and 80%, for the R91K, D, and P mutants, respectively. Relative stabilities of the wild type and R91 mutant proteins were examined using both thermal NADH:FR inactivation profiles and loss of intrinsic flavin fluorescence emission quenching. The results of these stability studies are shown in Fig. 5. For the wild-type domain, the loss of NADH:FR activity was 50% complete at a temperature, T m, of approximately 55°C. Examination of the various mutants, indicated that the R91K, A, and Q mutants exhibited the greatest stability with NADH:FR T ms of approximately 52–54°C, while the R91P mutant exhibited the greatest instability, corresponding to a T m of 42°C. Fluorescence emis-

228

MAROHNIC AND BARBER

FIG. 4. Fluorescence excitation and emission spectra of cb5r and R91 mutants. Excitation and emission spectra for samples of the oxidized H 4-tagged wild-type and selected R91 mutants, all at 5 ␮M FAD concentration, were recorded as described under Materials and Methods. Excitation spectra, recorded from 260 –510 nm and emission spectra, recorded from 460 – 610 nm are shown for the native enzyme samples and for the corresponding free FAD released following thermal denaturation of each protein. Individual spectra correspond to wild-type (—); R91K (. . .); R91A (- - - -), and R91P (- 䡠 - 䡠 -).

sion spectral measurements yielded very similar results for FAD release. For the wild-type domain, the midpoint in the flavin fluorescence enhancement profile corresponded to approximately 57°C, while this temperature was reduced for the R91K, A, and Q mutants to approximately 53–54°C. As demonstrated for the NADH:FR activity profiles, the R91P mutant exhibited the lowest temperature for the flavin fluorescence quenching profile, corresponding to approximately 42°C. The results of initial rate kinetic studies of the various cb5r R91 mutants, under conditions of constant pH (7.0) and ionic strength (␮ ⫽ 0.05) are shown in Table II. When compared to the wild-type domain, R91K retained the highest level of NADH:FR activity with a V max of 35 ␮mol NADH consumed/min/nmol FAD and corresponding to 70% of normal activity. The K m for NADH was also increased, to 10 ␮M, compared to the wild-type domain, suggesting that R91 substitution reduced the level of diaphorase activity. All the remaining FAD-containing mutants exhibited reduced NADH:FR activities in the order R91K ⬎ R91Q ⬎ R91D ⬎ R91A ⬎ R91L ⫽ R91P. The R91P mutant retained only 34% of the wild-type NADH:FR activity, corresponding to a V max of 17 ␮mol NADH consumed/ min/nmol FAD, while exhibiting an 18-fold increase in the K m for NADH to 105 ␮M. Chemical modification of arginine residues. To confirm that modification of arginine residues in cb5r resulted in decreased NADH:FR activity, chemical modification studies using both phenylglyoxal and 2,3-butandione were performed. Both reagents have been shown to modify the side chain of arginine residues

FIG. 5. Effects of temperature on the stabilities of cb5r and the R91 mutants. Stock solutions of H 4-tagged wild-type and R91 mutant domains (18 to 20 ␮M FAD) were heated in microfuge tubes as described under Materials and Methods. (Top) NADH:FR activities were assayed for all samples immediately following both dilution and at the conclusion of the inactivation profile. The average of the two values is plotted relative to the initial activity at 0°C. (Bottom) Flavin fluorescence emission spectra were determined at each temperature point and the emission maxima at 520 nm plotted as a percentage of total fluorescence. The total flavin fluorescence was obtained by measuring a similar dilution of the remaining stock solution following heating at 100°C for 30 min. Wild-type domain (E), R91K (䊐), R91A (Œ), R91Q (■), R91D (F), R91L (ƒ), R91P ({).

with a high degree of specificity (34, 35). Incubation of cb5r in the presence of phenylglyoxal resulted in both a time- and concentration-dependent inactivation of

TABLE II

Initial-Rate Kinetic Constants Obtained for the Various cb5r R91 Mutants NADH:FR

NADH:BR

Mutant

V max (units)

K mNADH (␮M)

K mFeCN6 (␮M)

V max (units)

K mcytb (␮M)

R91K R91A R91L R91P R91D R91Q R91H WT

35 23 17 17 26 28 ND 50

10 183 72 105 694 32 ND 6

5 8 6 5 8 9 ND 7

8 1 1 1 1 3 ND 17

10 15 12 15 13 15 ND 13

Note. ND, not determined.

CYTOCHROME b 5 REDUCTASE MUTAGENESIS

FIG. 6. Phenylglyoxal inactivation of cb5r and the R91A mutant. Samples of H 4-tagged cb5r (3 ␮M FAD) and R91A (4 ␮M FAD) were incubated with varying concentrations of phenylglyoxal, in both the absence and the presence of NADH (2 mM), in 100 mM bicarbonate buffer, containing 0.1 mM EDTA, pH 8.1, at 23°C. Aliquots were withdrawn at the indicated time intervals and assayed for NADH:FR activity, expressed as the percentage activity remaining with respect to a control sample maintained at 0°C in the absence of phenylglyoxal. (Top) cb5r incubated in the absence of phenylglyoxal (Œ), cb5r incubated in the presence of 1 mM (■), 3 mM (F), and 10 mM () phenylglyoxal and 3 mM phenylglyoxal and 2 mM NAD ⫹ (E). (Bottom) R91A incubated in the presence of 3 mM phenylglyoxal (F) and 3 mM phenylglyoxal and 2 mM NAD ⫹ (■).

NADH:FR activity, as shown in Fig. 6. In the presence of the highest concentration of phenylglyoxal examined, corresponding to 10 mM, 50% loss of activity was observed following 2 min incubation. Extended incubation periods, greater than 5 min, indicated that loss of activity reached a plateau corresponding to retention of approximately 40% of the initial NADH:FR activity. Incubation of cb5r with phenylglyoxal in the presence of NAD ⫹ provided significant protection of the enzyme against inactivation. A similar time- and concentration-dependent inactivation of cb5r was observed when phenylglyoxal was replaced by 2,3-butanedione. To investigate if R91 was the only arginine residue required for NADH:FR functionality, the R91A mutant was incubated with phenylglyoxal (3 mM) under the same conditions used for the native domain. Phenylglyoxal modification was observed to result in a similar, although decreased, time-dependent inactivation of NADH:FR activity, reaching a plateau of 75% activity remaining following 30 min incubation. Inclusion of NAD ⫹ in the incubation cocktail resulted in significant

229

protection of the R91A mutant against inactivation, corresponding to a loss of only 6% of the initial activity following 50 min incubation. To examine the spectroscopic and kinetic properties of the phenylglyoxal-modified form of the enzyme, cb5r was incubated with phenylglyoxal until only 40% of the initial NADH:FR activity remained. Further modification was terminated by the addition of excess L-arginine and the enzyme isolated by size exclusion FPLC. The UV/visible spectrum of the phenylglyoxal-modified enzyme exhibited absorption maxima at 271, 386, and 460 nm, comparable to the spectra obtained for the wild-type domain and suggesting that chemical modification of arginine residues did not result in any significant perturbation of the flavin environment. However, in contrast, initial-rate kinetic constants obtained for the NADH:FR activity of the phenylglyoxalmodified form of the enzyme indicated a decreased V max, corresponding to 5 ␮mol NADH/min/nmol FAD, and an increased K mNADH, corresponding to 120 ␮M, respectively, which were substantially altered from the corresponding values obtained for the wild-type domain (V max ⫽ 50 ␮mol NADH/min/nmol FAD, K mNADH ⫽ 6 ␮M). DISCUSSION

In this report, we have used site-directed mutagenesis, to examine the role of R91 in binding the FAD prosthetic group in cb5r. In the mature cb5r amino acid sequence, R91 corresponds to the initial residue in a seven-member segment of the enzyme’s primary sequence that has been identified to be highly conserved in the FNR family of flavoprotein oxidoreductases (24). A multiple sequence alignment of the corresponding seven residue segments and flanking amino acid residues from a number of constituents of the FNR family of flavoproteins in shown in Fig. 7. Within this grouping, a dominant consensus sequence, corresponding to the motif “RxY STxx NS” is readily identifiable, suggesting an important role in cb5r functionality that is potentially shared with other members of the FNR family of flavoprotein oxidoreductases. Within the FNR family of flavoproteins, X-ray structures are available for several members including spinach FNR (16), Burkholdetia cepaccia PDR (23), the flavin domains of corn (36) and spinach (37) assimilatory nitrate reductases, and the porcine (26) and rat (27) variants of cb5r. All five structures show similar domain architectures consisting of an amino-terminal flavin-binding domain and a carboxyl-terminal pyridine nucleotide-binding domain, with the flavin-binding domains exhibiting a six-stranded antiparallel ␤-barrel motif.

230

MAROHNIC AND BARBER

FIG. 7. Multiple sequence alignment of the residues comprising the “RxY STxx NS” putative FAD-binding motif in members of the FNR family of flavoprotein oxidoreductases. Amino acid sequences (44) comprising either the full-length or FAD-containing domains of selected members of the FNR family of flavoproteins were aligned using the “ClustalW” algorithm. The amino acid sequences correspond to rat cytochrome b 5 reductase (7), spinach nitrate reductase (20), M. capsulatus methane monooxygenase flavin subunit (22), P. putida 2-oxo-1,2-dihydroquinoline 8-monooxygenase (45), Acinetobacter sp. dimethyl sulfoxide oxygenase (46), Pseudomonas sp. TW3 4-nitrotoluene monooxygenase, reductase component (47), P. aeruginosa o-halobenzoate dioxygenase (48), Y. pseudotuberculosis CDP-6-deoxy-delta-3,4-glucoseen reductase (49), V. cholerae NQR enzyme complex, ␤ subunit (50), N. meningitidis NADH-ubiquinone reductase, subunit F (51), E. coli nitric oxide dioxygenase (52), spinach FNR (15), human nitric oxide synthetase (18), E. coli sulfite reductase (21), S. cerevisiae cytochrome P450 reductase (17), E. coli methionine synthase reductase (19), B. ceppacia phthalate dioxygenase reductase (23), A. calcoaceticus vanillate o-demethylase oxidoreductase (53), Acinetobacter sp. aniline dioxygenase, reductase component (54), E. coli ferredoxin reductase, electron transfer component (55), E. coli flavodoxin reductase (56), and human monoamine oxidase B (57). Residues comprising conserved amino acids in the “RxY STxx NS” motif are shown in bold italics. Superscripts indicate the positions of the starting amino acids in the corresponding primary sequences.

The X-ray structures of both porcine (26) and rat (27) cb5r indicate that R91, the initial residue of the “RxY STxx NS” consensus motif, is situated at the end of a ␤ strand (␤4) within the flavin-binding subdomain and is oriented toward the si-face of the FAD, on the opposite side of the flavin prosthetic group from the NADHbinding subdomain which opposes the re-face of the FAD. The R91 side-chain carbon atoms are positioned parallel to the FMN-ribose portion of the prosthetic group with the guanidinium moiety within hydrogenbonding distance (2.5–2.6 Å) of the pyrophosphate moiety. The rat cb5r structure indicates that both the ⑀ and the ␩2 nitrogens are involved in hydrogen bonding with oxygen atoms of both the FMN and AMP phosphate groups, respectively. Arginine 91 appears to make no additional contacts with the flavin prosthetic group. We have succeeded in generating a series of R91 mutants that include both conservative (K and H) and nonconservative (A, L, Q, D, and P) amino acid substi-

tutions. Of the latter substitutions, R91 has been successively replaced by aliphatic nonpolar (A, L, and P), polar (Q), and negatively charged (D) amino acid residues. All of these mutants have been purified to homogeneity and their spectroscopic and kinetic properties examined. The spectroscopic results obtained from the series of cb5r R91 mutants generated, confirm the role of this residue in assisting the binding of the FAD prosthetic group. As anticipated, all the mutants, with the exception of R91H, contained FAD, demonstrating that the majority of the substitutions of R91 did not preclude FAD incorporation and also had no influence on the FAD/FMN specificity of the domain, supporting the observation that flavin specificity in the FNR family of flavoproteins may be primarily controlled by a fourresidue motif that corresponds to the sequence 79RGGS in PDR (24), a FMN-containing flavoprotein and 124 GKMS in cb5r. However, it should be noted that not all the mutants generated were produced as flavopro-

CYTOCHROME b 5 REDUCTASE MUTAGENESIS

teins. R91H was the only mutant that was directly obtained as an apoprotein. The lack of flavin incorporation in this mutant may have been due to the instability of the protein since this mutant was obtained in very low yield following purification or may suggest a substantially altered secondary and/or tertiary structure that did not support FAD inclusion. For the remaining mutants, corresponding to R91K, A, L, Q, D, and P, the absence of any significant changes in the mutant far-UV CD spectra indicated that the amino acid substitutions did not alter the overall folding patterns of the mutant proteins compared to the wild-type domain. In contrast, perturbations of the visible absorption and CD spectra when compared to the wild-type domain suggested that with the exception of the R91K mutant, the remaining variants contained FAD bound in an altered environment or conformation. Spectroscopic studies of free flavin have indicated that red shifts in the visible absorbance maxima reflect a decrease in the hydrophobicity of the flavin, suggesting that substitutions of A, L, Q, D, and P for R91 resulted in a small increase in the polarity of the flavin environment (38). However, perturbations of the visible CD spectra for the majority of the mutants were significantly more dramatic. The visible CD spectra of the R91 mutants indicated that at least three optically active transitions occurred which were centered at approximately 305, 375, and 460 nm, respectively. The transition centered at 460 nm corresponds to a ␲–␲* transition (39) that was partially resolved into three vibronic bands in the wild-type domain and all six mutants, as previously observed in other flavoproteins (40). The transition at 375 nm is also a ␲–␲* transition with some n– ␲ * character (41). In the R91A, L, Q, D, and P mutants, while these CD bands showed no appreciable shifts in their wavelength maxima or spectral resolution, suggesting the absence of any significant changes in the hydrophobicity of the flavin environment, the CD transitions exhibited dramatic reversals of polarity when compared to either the wildtype domain or R91K mutant as well as a reduction in intensity for the 460-nm transition. The reversal in sign of these CD bands are due to the distortions in the flavin chromophor introduced by interaction with the protein side chains inducing opposite polarization in the flavin transitions. Similar differences in flavin visible CD have been observed for Clostridial flavodoxin and ferredoxin:NADP ⫹ reductase (39). Perturbations of the FAD environment were further supported by decreased quenching of the intrinsic flavin fluorescence and decreased thermal stability of the mutants. These results all suggest an altered conformation of the FAD chromophor in the mutant proteins. The results of the R91 site-directed mutagenesis studies are consistent with the cb5r X-ray structural data (26, 27) and the proposed role of this residue in

231

which the arginine side chain interacts with both the flavin FMN- and ADP-phosphate residues to form hydrogen bonds that effectively stabilize FAD incorporation. Replacement of R91 by K results in the loss of at least one of these hydrogen bonds, decreasing the interaction between the protein and FAD. For the remaining R91 mutants, the nonconservative substitutions result in complete loss of the hydrogen bond donors toward the FAD pyrophosphate moiety. These changes in FAD coordination are reflected in the altered visible absorption, CD and intrinsic fluorescence spectra of the mutants when compared to the wild-type enzyme. The visible CD spectra indicate that with the more radical residue substitutions, the spectra approach that observed for free FAD, which adopts a more linear conformation in solution, suggesting that the loss of H-bonding may result in a displacement of the FAD prosthetic group into the cleft between the FAD- and NADH-binding subdomains. This displacement toward the nucleotide-binding site would be expected to interfere with NADH-binding via a steric blockade effect and result in decreased diaphorase activities. An unanticipated result of these mutagenesis studies was that examination of the catalytic properties of the R91 cb5r mutants showed that all the amino acid substitutions resulted in proteins both with decreased NADH:FR and NADH:BR activities, supporting the conclusion that altered FAD binding also perturbed NADH utilization through a displacement into the substrate-binding cleft. This displacement would be anticipated to both reduce the enzyme’s substrate affinity and potentially alter the orientation between the bound NADH and FAD, reducing the efficiency of electron transfer. The initial-rate studies of the R91 mutants support this interpretation and revealed that both V max and K m values were affected, reflecting changes in the affinities of the various mutants for the reducing substrate, NADH, and reduced rates of electron transfer to FAD. In contrast, values obtained for the K m s for cytochrome b 5 for the individual mutants were unchanged from those of the wild-type domain, suggesting that R91 substitutions had little or no effect on the affinity of the cytochrome b 5 binding site. The results of the chemical modification studies using the arginine-specific reagents, phenylglyoxal (34) and 2,3-butanedione (42) were in good agreement with the results obtained using site-directed mutagenesis. Phenylglyoxal modification of the wild-type domain was observed to result in substantial irreversible inactivation of NADH:FR activity, while significant protection against loss of activity occurred in the presence of the competitive inhibitor, NAD ⫹. These results suggest that one or more arginine residues that are important for NADH:FR functionality are present in the vicinity of the NADH-binding site or at the interface of the

232

MAROHNIC AND BARBER

NADH- and FAD-binding domains. Additional chemical modification studies of the R91A mutant demonstrated that substitution of R91 by A afforded significant protection against phenylglyoxal inactivation with retention of protection by NAD ⫹. The combination of these results with those obtained for the wild-type domain suggest that while phenylglyoxal modification of R91 results in decreased NADH:FR activity, additional arginine residue(s) may also be involved in the interaction with NADH. The results of our studies of the role of R91 in cb5r can be compared with those published for the application of site-directed mutagenesis to examine the function of the corresponding arginine residue (R42) in the related flavoprotein, human monoamine oxidase B (MAO B), which also contains the RxY STxx NS flavin-binding motif (43). Substitution of R42 with A in human MAO B, resulted in the production of the corresponding apoprotein that was devoid of both FAD incorporation and catalytic activity. In contrast, the R42K mutation resulted in production of a protein that retained 14% of wild-type enzyme activity and exhibited decreased flavin incorporation. These results suggest that substitutions of R42 in MOA B are tolerated to a much lower degree than the corresponding substitutions in cb5r. In contrast to cb5r, which contains a non-covalent FAD prosthetic group, MOA B contains covalently bound FAD attached via an 8 ␣-S-cysteinyl linkage. In MAO B, R42 has been proposed to initially bind the FAD to the protein through noncovalent hydrogen-bonding interactions prior to formation of the final 8S-cysteinyl covalent linkage. The results obtained for cb5r indicate that a number of alternative residues can be substituted for R91 with retention of both FAD binding and significant NADH:FR activity, although an altered cofactor architecture and reduced catalytic activity were apparent. Thus, while our results have suggested that R91 is an important residue in enhancing cb5r functionality, it is not essential for FAD incorporation. ACKNOWLEDGMENTS This work was supported by grants GM 32696 from the National Institutes of Health and 9701708 and 9910034V from the American Heart Association, Florida/Puerto Rico Affiliate.

REFERENCES 1. Spatz, L., and Strittmatter, P. (1973) J. Biol. Chem. 248, 793– 799. 2. Iyanagi, T., Watanabe, S., and Anan, K. F. (1984) Biochemistry 23, 1418 –1425. 3. Oshino, N., Imai, Y., and Sato, R. (1971) J. Biochem. (Tokyo) 69, 155–167. 4. Keyes, S. R., and Cinti, D. L. (1980) J. Biol. Chem. 255, 11,357– 11,364.

5. Reddy, V. V. R., Kupfer, D., and Capsi, E. (1977) J. Biol. Chem. 252, 2797–2801. 6. Hildebrant, A., and Estabrook, R. W. (1971) Arch Biochem. Biophys. 143, 66 –79. 7. Pietrini, G., Carrera, P., and Borgese, N. (1988) Proc. Natl. Acad. Sci. USA 85, 7246 –7250. 8. Kitao, T., Sugita, Y., Yoneyama, Y., and Hattori, K. (1974) Blood. 44, 879 – 884. 9. Murakami, K., Yubisui, T., Takeshita, M., and Miyata, T. (1989) J. Biochem. (Tokyo) 105, 312–317. 10. Ozols, J., Carr, S. A., and Strittmatter, P. (1984) J. Biol. Chem. 259, 13349 –13354. 11. Iyanagi, T. (1977) Biochemistry 16, 2725–2730. 12. Crankshaw, D. L., Crankshaw, C. L., and Husby, A. D. (1987) BioChromatography 2, 196 –204. 13. Ozols, J., Korza, G., Heinemann, F., Hediger, M., and Strittmatter, P. (1985) J. Biol. Chem. 260, 11953–11961. 14. Strittmatter, P. (1962) J. Biol. Chem. 237, 3250 –3254. 15. Strittmatter, P. (1965) J. Biol. Chem. 240, 4481– 4487. 16. Karplus, P. A., Daniels, M. J., and Herriott, J. R. (1991) Science 251, 60 – 65. 17. Yabusaki, Y., Murakami, H., and Ohkawa, H. (1988) J. Biochem. 103, 1004 –1010. 18. Geller, D. A., Lowenstein, C. J., Shapiro, R. A., Nussler, A. K., DiSilvio, M., Wang, S. C., Nakayama, D. K., Simmons, R. L., Snyder, S. H., and Billiar, T. R. (1993) Proc. Natl. Acad. Sci. USA 90, 3491–3495. 19. Leclerc, D., Odievre, M.-H., Wu, Q., Wilson, A., Huizenga, J. J., Rozen, R., Scherer, S. W., and Gravel, R. A. (1999) Gene 240, 75– 88. 20. Prosser, I. M., and Lazarus, C. M. (1990) Plant Mol. Biol. 15, 187–190. 21. Ostrowski, J., Barber, M. J., Rueger, D. C., Miller, B. E., Siegel, L. M., and Kredich, N. M. (1989) J. Biol. Chem. 264, 15,796 – 15,808. 22. Stainthorpe, A. C., Lees, V., Salmond, G. P. C., Dalton, H., and Murrell, J. C. (1990) Gene 91, 27–34. 23. Correll, C. C., Batie, C. J., Ballou, D. P., and Ludwig, M. L. (1992) Science 258, 1604 –1610. 24. Correll, C. C., Ludwig, M. L., Bruns, C. M., and Karplus, P. A. (1993) Protein Sci. 2, 2112–2133. 25. Marohnic, C. C., and Barber, M. J. (1999) FASEB J. 13, A1437. 26. Nishida, H., Inaka, K., Yamanaka, M., Kaido, S., Kobayashi, K., and Miki, K. (1995) Biochemistry 34, 2763–2767. 27. Marohnic, C. C., Bewley, M., and Barber, M. J. (2001) FASEB J., 15, A525. 28. Barber, M. J., and Quinn, G. B. (1996) Protein Expression Purif. 7, 41– 47. 29. Beck-von Bodman, S. B., Schuler, M. A., Jollie, D. R., and Sligar, S. G. (1986) Proc. Natl. Acad. Sci. USA 83, 9443–9447. 30. Laemmli, U. K. (1970) Nature 227, 680 – 685. 31. Chen, G. C., and Yang, J. T. (1977) Anal. Lett. 10, 1195–1207. 32. Kay, C. J., and Barber, M. J. (1986) J. Biol. Chem. 261, 14,125– 14,129. 33. Trimboli, A. J., Quinn, G. B., Smith, E. T., and Barber, M. J. (1996) Arch. Biochem. Biophys. 331, 117–126. 34. Berghauser, J., and Shirmer, R. H. (1978) Biochim. Biophys. Acta. 537, 428 – 435. 35. Nagai, K., and Thogerson, H. C. (1984) Nature 309, 810 – 812. 36. Lu, G., Lindqvist, Y., Schneider, G., Dwivedi, U., and Campbell, W. (1995) J. Mol. Biol. 248, 931–948.

CYTOCHROME b 5 REDUCTASE MUTAGENESIS 37. Bewley, M., and Barber, M. J. (2001) FASEB J. 15, A525. 38. Harbury, H. A., La Nove, K. F., Loach, P. A., and Amick, R. M. (1959) Proc. Natl. Acad. Sci. USA 45, 1708 –1717. 39. Edmondoson, D. E., and Tollin, G. (1971) Biochemistry 10, 113– 123. 40. Penzer, G. R., and Radda, G. K. (1966) Q. Rev. Chem. Soc. 21, 43– 67. 41. Eweg, J. D., Muller, F., van Dam, H., Terpstra, A., and Oskam, A. (1980) J. Am. Chem. Soc. 102, 51– 61. 42. Vallejos, R. H., Lescano, W. I. M., and Lucero, H. A. (1978) Arch. Biochem. Biophys. 190, 578 –584. 43. Kirksey, T. J., Kwan, S-W., and Abell, C. W. (1998) Biochemistry 37, 12,360 –12,366. 44. Benson, D. A., Karsch-Mizrachi, I., Lipman, D. J., Ostell, J., Rapp, B. A., and Wheeler, D. L. (2000) Genbank. Nucleic Acids Res. 28, 15–18. 45. Rosche, B., Tshisuaka, B., Hauer, B., Lingens, F., and Fetzner, S. (1997) J. Bacteriol. 179, 3549 –3554. 46. Horinouchi, M., Kasuga, K., Nojiri, H., Yamane, H., and Omori, T. (1997) FEMS Microbiol. Lett. 155, 99 –105. 47. James, K. D., and Williams, P. A. (1998) J. Bacteriol. 180, 2043– 2049. 48. Arment, A. R., Yuroff, A. S., Perez-Lesher, J., Xia, X., and Hickey, W. J. (1998) o-halobenzoate dioxygenase GenBank AAC69483.

233

49. Skurnik, M., Peippo, A., and Ervela, E. (1999) CDP-6-deoxydelta-3,4-glucoseen reductase GenBank CAB63289. 50. Hase, C. C., and Mekalanos, J. J. (1999) Proc. Natl. Acad. Sci. USA 96, 3183–3187. 51. Parkhill, J., Achtman, M., James, K. D., Bentley, S. D., Churcher, C., Klee, S. R., Morelli, G., Basham, D., Brown, D., Chillingworth, T., Davies, R. M., Davis, P., Devlin, K., Feltwell, T., Hamlin, N., Holroyd, S., Jagels, K., Leather, S., Moule, S., Mungall, K., Quail, M. A., Rajandream, M. A., Rutherford, K. M., Simmonds, M., Skelton, J., Whitehead, S., Spratt, B. G., and Barrell, B. G. (2000) Nature 404, 502–506. 52. Vasudevan, S. G., Armarego, W. L., Shaw, D. C., Lilley, P. E., Dixon, N. E., and Poole, R. K. (1991) Mol. Gen. Genet. 226, 49 –58. 53. Segura, A., and Ornston, N. L. (1999) vanillate o-demethylase oxidoreductase GenBank O24840. 54. Takeo, M., Fujii, T., and Maeda, Y. (1998) J. Ferment. Bioeng. 85, 17–24. 55. Ferrandez, A., Prieto, M. A., Garcia, J. L., and Diaz, E. (1997) FEBS Lett. 406, 23–27. 56. Bianchi, V., Reichard, P., Eliasson, R., Pontis, E., Krook, M., Jornvall, H., and Haggard-Ljungquist, E. (1993) J. Bacteriol. 175, 1590 –1595. 57. Bach, A. W., Lan, N. C., Johnson, D. L., Abell, C. W., Bembenek, M. E., Kwan, S. W., Seeburg, P. H., and Shih, J. C. (1988) Proc. Natl. Acad. Sci. USA 85, 4934 – 4938.