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• W1996652032 abstract "The oxygenase domain of inducible nitric-oxide synthase exists as a functional tight homodimer in the presence of the substrate l-arginine and the cofactor tetrahydrobiopterin (H4B). In the absence of H4B, the enzyme is a mixture of monomer and loose dimer. We show that exposure of H4B-free enzyme to NO induces dissociation of the loose dimer into monomers in a reaction that follows single exponential decay kinetics with a lifetime of ∼300 min. It is followed by a faster autoreduction reaction of the heme iron with a lifetime of ∼30 min and the concurrent breakage of the proximal iron-thiolate bond, forming a five-coordinate NO-bound ferrous species. Mass spectrometry revealed that the NO-induced monomerization is associated with intramolecular disulfide bond formation between Cys104 and Cys109, located in the zinc-binding motif. The regulatory effect of NO as a dimer inhibitor is discussed in the context of the structure/function relationships of this enzyme. The oxygenase domain of inducible nitric-oxide synthase exists as a functional tight homodimer in the presence of the substrate l-arginine and the cofactor tetrahydrobiopterin (H4B). In the absence of H4B, the enzyme is a mixture of monomer and loose dimer. We show that exposure of H4B-free enzyme to NO induces dissociation of the loose dimer into monomers in a reaction that follows single exponential decay kinetics with a lifetime of ∼300 min. It is followed by a faster autoreduction reaction of the heme iron with a lifetime of ∼30 min and the concurrent breakage of the proximal iron-thiolate bond, forming a five-coordinate NO-bound ferrous species. Mass spectrometry revealed that the NO-induced monomerization is associated with intramolecular disulfide bond formation between Cys104 and Cys109, located in the zinc-binding motif. The regulatory effect of NO as a dimer inhibitor is discussed in the context of the structure/function relationships of this enzyme. Nitric-oxide synthase (NOS) 4The abbreviations used are: NOS, nitric-oxide synthase; iNOS, inducible nitric-oxide synthase; H4B, tetrahydrobiopterin; NOSoxy, nitric-oxide synthase oxygenase domain; EPPS, 4-(2-hydroxyethyl)-1-piperazinepropanesulfonic acid; PAR, 4-(2-pyridylazo)resorcinol monosodium salt; 6C, six-coordinate; 5C, five-coordinate. 4The abbreviations used are: NOS, nitric-oxide synthase; iNOS, inducible nitric-oxide synthase; H4B, tetrahydrobiopterin; NOSoxy, nitric-oxide synthase oxygenase domain; EPPS, 4-(2-hydroxyethyl)-1-piperazinepropanesulfonic acid; PAR, 4-(2-pyridylazo)resorcinol monosodium salt; 6C, six-coordinate; 5C, five-coordinate. catalyzes the formation of NO from oxygen and l-Arg via a consecutive two-step reaction using NADPH as the electron source (1Stuehr D.J. Biochim. Biophys. Acta. 1999; 1411: 217-230Crossref PubMed Scopus (798) Google Scholar, 2Alderton W.K. Cooper C.E. Knowles R.G. Biochem. J. 2001; 357: 593-615Crossref PubMed Scopus (3185) Google Scholar, 3Marletta M.A. Adv. Exp. Med. Biol. 1993; 338: 281-284Crossref PubMed Google Scholar). In the first step of the reaction, l-Arg is hydroxylated to N-hydroxyarginine; and in the second step, N-hydroxyarginine is oxidized to citrulline and NO. The three major isoforms, inducible NOS (iNOS), endothelial NOS, and neuronal NOS (found in macrophages, endothelial cells, and neuronal tissues, respectively), produce NO that functions as a cytotoxic agent, a vasodilator, and a neurotransmitter, respectively (4Nathan C. Xie Q.W. Cell. 1994; 78: 915-918Abstract Full Text PDF PubMed Scopus (2723) Google Scholar). The homodimeric enzyme consists of a reductase domain, which binds FMN, FAD, and NADPH, and an oxygenase domain, which binds the heme and tetrahydrobiopterin (H4B) cofactors. During catalysis, electrons flow from NADPH through FMN and FAD in the reductase domain of one subunit of the homodimer to the oxygenase domain of the other subunit (5Siddhanta U. Presta A. Fan B. Wolan D. Rousseau D.L. Stuehr D.J. J. Biol. Chem. 1998; 273: 18950-18958Abstract Full Text Full Text PDF PubMed Scopus (181) Google Scholar, 6Panda K. Ghosh S. Stuehr D.J. J. Biol. Chem. 2001; 276: 23349-23356Abstract Full Text Full Text PDF PubMed Scopus (109) Google Scholar). The crystal structures of the oxygenase domain of all three isoforms have been determined (7Crane B.R. Arvai A.S. Ghosh D.K. Wu C. Getzoff E.D. Stuehr D.J. Tainer J.A. Science. 1998; 279: 2121-2126Crossref PubMed Scopus (619) Google Scholar, 8Raman C.S. Li H. Martasek P. Kral V. Masters B.S. Poulos T.L. Cell. 1998; 95: 939-950Abstract Full Text Full Text PDF PubMed Scopus (568) Google Scholar, 9Li H. Raman C.S. Glaser C.B. Blasko E. Young T.A. Parkinson J.F. Whitlow M. Poulos T.L. J. Biol. Chem. 1999; 274: 21276-21284Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar, 10Fischmann T.O. Hruza A. Niu X.D. Fossetta J.D. Lunn C.A. Dolphin E. Prongay A.J. Reichert P. Lundell D.J. Narula S.K. Weber P.C. Nat. Struct. Biol. 1999; 6: 233-242Crossref PubMed Scopus (395) Google Scholar). They show that the heme is coordinated by a cysteine residue on the proximal side, as in cytochrome P450-type enzymes, and that the substrate (l-Arg or N-hydroxyarginine) binds above the heme iron atom in the distal pocket, whereas the cofactor (H4B) binds along the side of the heme. It is well accepted that dimerization is essential for NOS function (1Stuehr D.J. Biochim. Biophys. Acta. 1999; 1411: 217-230Crossref PubMed Scopus (798) Google Scholar, 11Marletta M.A. Cell. 1994; 78: 927-930Abstract Full Text PDF PubMed Scopus (805) Google Scholar). The heme group, the H4B cofactor, and the substrate have all been shown to contribute to dimer stability (12Klatt P. Schmidt K. Lehner D. Glatter O. Bachinger H.P. Mayer B. EMBO J. 1995; 14: 3687-3695Crossref PubMed Scopus (263) Google Scholar, 13Klatt P. Pfeiffer S. List B.M. Lehner D. Glatter O. Bachinger H.P. Werner E.R. Schmidt K. Mayer B. J. Biol. Chem. 1996; 271: 7336-7342Abstract Full Text Full Text PDF PubMed Scopus (166) Google Scholar, 14Mayer B. Wu C. Gorren A.C. Pfeiffer S. Schmidt K. Clark P. Stuehr D.J. Werner E.R. Biochemistry. 1997; 36: 8422-8427Crossref PubMed Scopus (98) Google Scholar, 15Panda K. Rosenfeld R.J. Ghosh S. Meade A.L. Getzoff E.D. Stuehr D.J. J. Biol. Chem. 2002; 277: 31020-31030Abstract Full Text Full Text PDF PubMed Scopus (106) Google Scholar, 16Chen Y. Panda K. Stuehr D.J. Biochemistry. 2002; 41: 4618-4625Crossref PubMed Scopus (32) Google Scholar). In iNOS, the N-terminal region (between residues 76 and 111, comprising a β-hairpin hook and a CXXXC zinc-binding motif) is also believed to be important for stabilizing the dimeric structure. Crane et al. (17Crane B.R. Rosenfeld R.J. Arvai A.S. Ghosh D.K. Ghosh S. Tainer J.A. Stuehr D.J. Getzoff E.D. EMBO J. 1999; 18: 6271-6281Crossref PubMed Scopus (97) Google Scholar) reported that the N-terminal region of the iNOS oxygenase domain (iNOSoxy) can be in either a “swapped” or an “unswapped” conformation, as illustrated in Fig. 1. In the unswapped conformation, Cys104 and Cys109 in the zinc-binding motif of each subunit of the dimer are tetrahedrally coordinated to a single zinc ion at the dimer interface, and the β-hairpin hook interacts primarily with its own subunit; in the swapped conformation, Cys109 forms a self-symmetric disulfide bond across the dimer interface, and the β-hairpin hook in one subunit of the dimer interacts primarily with the other subunit across the interface (17Crane B.R. Rosenfeld R.J. Arvai A.S. Ghosh D.K. Ghosh S. Tainer J.A. Stuehr D.J. Getzoff E.D. EMBO J. 1999; 18: 6271-6281Crossref PubMed Scopus (97) Google Scholar). Crane et al. proposed that the conformational switch between the two structures may play an important role in NOS stability and function in vivo. It has been found that NO produced from the catalytic reaction in iNOS not only can rebind to the heme iron, thereby directly inhibiting the turnover of the enzyme (18Abu-Soud H.M. Wang J. Rousseau D.L. Fukuto J.M. Ignarro L.J. Stuehr D.J. J. Biol. Chem. 1995; 270: 22997-23006Abstract Full Text Full Text PDF PubMed Scopus (199) Google Scholar, 19Abu-Soud H.M. Ichimori K. Nakazawa H. Stuehr D.J. Biochemistry. 2001; 40: 6876-6881Crossref PubMed Scopus (63) Google Scholar), but also can induce monomerization of the functional dimers (16Chen Y. Panda K. Stuehr D.J. Biochemistry. 2002; 41: 4618-4625Crossref PubMed Scopus (32) Google Scholar). Although binding of l-Arg and H4B to iNOS promotes the formation of a “tight” dimer, which is resistant to monomerization by NO, it has been shown that the NO-induced monomers cannot be reverted back to the dimeric state by the addition of l-Arg and H4B (16Chen Y. Panda K. Stuehr D.J. Biochemistry. 2002; 41: 4618-4625Crossref PubMed Scopus (32) Google Scholar). The dimer inhibition function of NO has also been reported in endothelial NOS by Ravi et al. (20Ravi K. Brennan L.A. Levic S. Ross P.A. Black S.M. Proc. Natl. Acad. Sci. U. S. A. 2004; 101: 2619-2624Crossref PubMed Scopus (195) Google Scholar), who discovered that exogenous NO induces S-nitrosylation of a Cys residue in the zinc-binding motif, thereby reducing the dimer level and the associated enzymatic activity. Another type of dimer inhibition function of NO in iNOS has been demonstrated in the RAW 264.7 mouse macrophage cell line by Albakri and Stuehr (21Albakri Q.A. Stuehr D.J. J. Biol. Chem. 1996; 271: 5414-5421Abstract Full Text PDF PubMed Scopus (139) Google Scholar), who found that NO produced by iNOS induced by cytokines limits the intracellular assembly of iNOS into the dimeric form by preventing heme insertion and decreasing heme availability. Although it is clear that NO plays an important role in regulating the monomer-dimer equilibrium, the molecular mechanism underlying the NO-induced structural transition remains poorly understood. Here, we systematically studied the interaction between NO and wild-type iNOSoxy as well as two mutants (D92A and K82A) by optical absorption and resonance Raman spectroscopies; in addition, the chemical modifications of the protein matrix induced by NO were examined by mass spectrometry. The data reveal a detailed mechanism of the inhibitory and regulatory effects of NO, as a heme iron ligand, a cysteine-modifying agent, and an inhibitor of dimerization. (6R)-5,6,7,8-Tetrahydro-l-biopterin was purchased from Alexis Biochemicals (San Diego, CA). All other reagents were from Sigma. Murine wild-type iNOSoxy and mutants were expressed in Escherichia coli, purified, and prepared as described previously (17Crane B.R. Rosenfeld R.J. Arvai A.S. Ghosh D.K. Ghosh S. Tainer J.A. Stuehr D.J. Getzoff E.D. EMBO J. 1999; 18: 6271-6281Crossref PubMed Scopus (97) Google Scholar, 22Li D. Stuehr D.J. Yeh S.-R. Rousseau D.L. J. Biol. Chem. 2004; 279: 26489-26499Abstract Full Text Full Text PDF PubMed Scopus (63) Google Scholar). For urea-containing samples, urea was added from a 12 m stock solution and allowed to equilibrate for 3 h prior to the measurements. To form the NO-bound complexes, 400 μl of 1 atm NO was injected into N2-purged solutions sealed in an optical cuvette. All spectroscopic measurements were made under anaerobic conditions. Optical Absorption and Resonance Raman Measurements—Optical absorption and resonance Raman spectra were obtained as described previously (22Li D. Stuehr D.J. Yeh S.-R. Rousseau D.L. J. Biol. Chem. 2004; 279: 26489-26499Abstract Full Text Full Text PDF PubMed Scopus (63) Google Scholar). For these measurements, the protein was kept in 40 mm EPPS at pH 7.6. The concentrations used are listed in the figure legends. The time-dependent optical spectra were deconvoluted using a program written with Mathcad software (Mathsoft, Cambridge, MA). In each case, the reference for the six-coordinate NO-bound species was taken immediately after the addition of NO. The reference spectrum for the five-coordinate species for all fittings was taken following incubation of the 4 m urea-treated sample with NO for >15 h. The kinetic traces were fitted using commercial software (Origin, RockWare, Golden, CO). Mass Spectrometric Measurements—The NO-treated iNOSoxy samples were generated by incubating the enzyme with NO for 12 h under anaerobic conditions at room temperature. The reaction was quenched by purging the NO with argon gas. All samples were then digested aerobically with modified trypsin (sequence-grade; Promega Corp.) in ammonium bicarbonate buffer overnight at 37 °C. All digestion products were desalted, separated by gradient elution with a Dionex reverse phase capillary/nano high pressure liquid chromatography system, and analyzed using an Applied Biosystems QSTAR XL tandem mass spectrometer with the hybrid quadrupole time-of-flight configuration. IDA (information-dependent acquisition) software was employed for automatic acquisition of mass spectrometric and tandem mass spectrometric data. The iNOSoxy sample without NO treatment was used as a control. To test the presence of disulfide bonds, half of the digestion products of the urea-treated sample were treated with 10 mm dithiothreitol in 0.1 m ammonium bicarbonate to reduce possible disulfide bonds; the free cysteine residues were then alkylated with freshly prepared iodoacetamide (55 mm in 0.1 m ammonium bicarbonate buffer); and the resulting sample was subsequently subjected to the mass spectrometric analysis. Size Exclusion Chromatographic Analysis—The Superdex 200 10/30 GL column was purchased from Amersham Biosciences. The iNOS samples (100 μl of 10-20 μm) were incubated first with 50 μm H4B and 5 mm l-Arg for >3 h and then with 0-7 m urea for 3 h. They were loaded onto the column pre-equilibrated with 40 mm EPPS at pH 7.6 at the specified concentrations of urea. The flow rate was 0.35 ml/min for all measurements, which were carried out at 4 °C, and the samples were run for 1.5 column volumes. The PAR Zinc Chelation Assay—4-(2-Pyridylazo)resorcinol monosodium salt (PAR) was purchased from Sigma. The zinc content was measured by the PAR assay as described previously (23Miller R.T. Martasek P. Raman C.S. Masters B.S. J. Biol. Chem. 1999; 274: 14537-14540Abstract Full Text Full Text PDF PubMed Scopus (33) Google Scholar) with slight modifications. The iNOSoxy·NO complex was prepared as described above. PAR was added to yield a final concentration of 8 μm. As a control, PAR was added to a ferric iNOSoxy sample at the same concentration. To isolate the contributions in the spectra from PAR in ferric iNOSoxy plus PAR and iNOSoxy·NO plus PAR, the corresponding iNOSoxy spectra were subtracted. To evaluate the effect of NO on the dimeric interactions in iNOSoxy, the substrate- and cofactor-free ferric enzyme was subjected to NO, and the reactions were monitored by optical absorption spectroscopy as a function of time. As shown in Fig. 2a, immediately after the addition of NO, a species with a Soret absorption maximum at 439 nm and visible absorption bands at 549 and 580 nm was produced. It was assigned as a six-coordinate (6C) NO-bound ferric iNOSoxy complex because its spectra are analogous to those of other reported 6C NO-bound NOS complexes (24Wang J. Rousseau D.L. Abu-Soud H.M. Stuehr D.J. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 10512-10516Crossref PubMed Scopus (125) Google Scholar). The 6C NO-bound ferric enzyme gradually converted to a species with a Soret maximum at ∼390 nm over an ∼300-min time period with a clear isosbestic point at 411 nm. The new species was assigned as a five-coordinate (5C) derivative of iNOSoxy because its spectral properties are similar to those of other 5C derivatives of NOS (22Li D. Stuehr D.J. Yeh S.-R. Rousseau D.L. J. Biol. Chem. 2004; 279: 26489-26499Abstract Full Text Full Text PDF PubMed Scopus (63) Google Scholar, 24Wang J. Rousseau D.L. Abu-Soud H.M. Stuehr D.J. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 10512-10516Crossref PubMed Scopus (125) Google Scholar). The properties of the 5C species are discussed below. To further evaluate the mechanism of the 6C-to-5C conversion, we deconvoluted each time-dependent spectrum into a linear combination of the spectrum of the 6C NO-bound ferric species and that of the 5C species. Typical examples demonstrating the reliability of the deconvolution process are shown in supplemental Fig. S1. The resulting population of the 5C species is plotted as a function of time in Fig. 2a (inset), and the associated kinetic trace was best fit with a double exponential function with lifetimes of 21 and 287 min. A similar reaction was observed for iNOSoxy in the presence of l-Arg as shown in Fig. 2b. Although the kinetic lifetimes (23 and 396 min) were only slightly altered upon the addition of l-Arg, the relative amplitude of the slow phase increased from 49% in the absence of l-Arg to 67% in its presence. The conversion of the 6C NO-bound ferric derivative to the 5C species was inhibited by the binding of H4B, either with or without l-Arg (supplemental Fig. S2). On the basis of gel filtration analysis, Panda and co-workers (15Panda K. Rosenfeld R.J. Ghosh S. Meade A.L. Getzoff E.D. Stuehr D.J. J. Biol. Chem. 2002; 277: 31020-31030Abstract Full Text Full Text PDF PubMed Scopus (106) Google Scholar) have reported that, in the absence of H4B, iNOSoxy is in equilibrium between a monomeric form and a “loose” dimeric form and that l-Arg binding shifts the equilibrium toward the loose dimer, whereas H4B binding generates a tight dimer. Accordingly, we postulated that the production of the 5C species in the reaction shown in Fig. 2 was a consequence of the lack of strong dimeric interactions in the absence of H4B. We attribute the slow phase to the reaction of the loose dimer because the amplitude of the slow phase increased from 49 to 67% when l-Arg was added, and we attribute the fast phase to the reaction of the monomeric fraction of the iNOSoxy samples because it decreased correspondingly in the presence of l-Arg. To test this hypothesis, we examined the reaction between the monomeric form of iNOSoxy and NO using a urea-induced monomer as a model. It has been reported that 5 m urea induces 100% conversion of the dimeric enzyme into monomers, but it is accompanied by significant loss of the heme group because of denaturation; on the other hand, reducing the urea concentration to 3 m can induce only ∼94% of the dimer to convert to its monomeric form (15Panda K. Rosenfeld R.J. Ghosh S. Meade A.L. Getzoff E.D. Stuehr D.J. J. Biol. Chem. 2002; 277: 31020-31030Abstract Full Text Full Text PDF PubMed Scopus (106) Google Scholar). To find the best conditions for generating the monomeric enzyme without denaturation, we titrated iNOSoxy with urea and found that 4 m urea was an optimum condition for generating the monomeric enzyme without heme loss (supplemental Fig. S3a). The monomeric state of the 4 m urea-treated iNOSoxy sample was confirmed by MALDI-TOF mass spectrometric measurements (data not shown) and by gel filtration analysis (supplemental Fig. S4). As shown in Fig. 3a, exposure of 4 m urea-treated iNOSoxy to NO instantaneously produced a 6C NO-bound ferric species with a Soret maximum at 439 nm, just as observed in the urea-free samples shown in Fig. 2; in addition, an analogous spectral transition from the 6C NO-bound ferric derivative to a 5C species was observed, although with altered kinetics. To gain quantitative information, the population of the 5C species was estimated by spectral deconvolution of the optical absorption data and was plotted as a function of reaction time in Fig. 3a (inset). The resulting kinetic trace was best fit with a single exponential function with a lifetime of ∼30 min. This lifetime is similar to that of the fast phase (21-23 min) obtained in the absence of urea (Fig. 2), consistent with the scenario that the fast phase originates from the monomeric enzyme. It should be noted that, in the gel filtration measurements in 4 m urea, a small amount of dimer was detected, but its elution volume was spread out compared with that in the absence of urea. We attribute it to some very loose dimer, which reacted with NO as rapidly as the monomer. To further confirm that the fast phase indeed originates from the monomeric derivative and to eliminate any possible side effects caused by the addition of urea, the NO reaction was examined using two iNOSoxy mutants (D92A and K82A) that adopt a pure monomeric conformation in the absence of H4B (25Ghosh D.K. Crane B.R. Ghosh S. Wolan D. Gachhui R. Crooks C. Presta A. Tainer J.A. Getzoff E.D. Stuehr D.J. EMBO J. 1999; 18: 6260-6270Crossref PubMed Scopus (62) Google Scholar). Fig. 3b shows the time-dependent optical absorption spectra of the D92A mutant of iNOSoxy following exposure to NO. Again, the instantaneously formed 6C NO-bound enzyme with a Soret band at 439 nm converted to the 5C species with a Soret maximum at 390 nm and a single exponential decay rate of ∼28 min, similar to that observed in the urea-stabilized monomeric wild-type enzyme sample. Similar kinetic behavior was observed with the K82A mutant (supplemental Fig. S3b), confirming that the 20-30-min kinetic phase originates from the monomeric form of the enzyme. On the basis of these data, we concluded that NO binding to the ferric protein in either the monomeric or dimeric state instantaneously produces a 6C NO-bound ferric derivative with indistinguishable optical absorption spectra with a Soret transition maximum at 439 nm. Subsequently, the fast phase is attributed to the transition from the 6C NO-bound state of the monomeric protein ([M-NO]6C) to the 5C species ([M]5C-NO), whereas the slow phase is ascribed to the same reaction originating from the loose dimer ([D-NO]6C) as described in Equation 1. [D-NO]6C→[M-NO]6C→[M]5C-NO(Eq. 1) Here, the formation of the 5C species is rate-limited by the monomerization of the dimer with an apparent lifetime of ∼300-400 min. To gain insights into the nature of the 5C species, the NO-treated samples were examined by resonance Raman spectroscopy. As shown in Fig. 4, the resonance Raman spectra of the 5C species with a Soret maximum at 390 nm generated in the presence and absence of urea are very similar (upper and middle traces), indicating that the two 5C species are the same. Because these spectra are virtually identical to those of 5C NO-bound ferrous derivatives of a variety of heme proteins as characterized by the heme modes located at 349, 677, and 756 cm-1 and a broad Fe-NO stretching mode (νFe-NO) in the 520-526 cm-1 region (26Deinum G. Stone J.R. Babcock G.T. Marletta M.A. Biochemistry. 1996; 35: 1540-1547Crossref PubMed Scopus (183) Google Scholar, 27Vogel K.M. Hu S. Spiro T.G. Dierks E.A. Yu A.E. Burstyn J.N. J. Biol. Inorg. Chem. 1999; 4: 804-813Crossref PubMed Scopus (45) Google Scholar, 28Andrew C.R. Green E.L. Lawson D.M. Eady R.R. Biochemistry. 2001; 40: 4115-4122Crossref PubMed Scopus (77) Google Scholar, 29Reynolds M.F. Parks R.B. Burstyn J.N. Shelver D. Thorsteinsson M.V. Kerby R.L. Roberts G.P. Vogel K.M. Spiro T.G. Biochemistry. 2000; 39: 388-396Crossref PubMed Scopus (89) Google Scholar, 30Lukat-Rodgers G.S. Rodgers K.R. Biochemistry. 1997; 36: 4178-4187Crossref PubMed Scopus (78) Google Scholar), the resonance Raman spectrum of the 5C NO-bound ferrous derivative of the iNOSoxy complex was also obtained (Fig. 4, lower trace). Here, the 5C NO-bound ferrous derivative was formed by directly adding NO to the substrate- and cofactor-free ferrous enzyme because the conversion of the 6C ferrous NO complex to its 5C form has been demonstrated previously in both iNOSoxy and neuronal NOSoxy by Stuehr and co-workers (31Abu-Soud H.M. Wu C. Ghosh D.K. Stuehr D.J. Biochemistry. 1998; 37: 3777-3786Crossref PubMed Scopus (111) Google Scholar, 32Huang L. Abu-Soud H.M. Hille R. Stuehr D.J. Biochemistry. 1999; 38: 1912-1920Crossref PubMed Scopus (47) Google Scholar). The small differences in the 378 and 524 cm-1 regions are attributed to differences in the contributions of the laser plasma lines. The identical features in the three traces shown in Fig. 4 indicate that exposure of the ferric derivative of H4B-free iNOSoxy to NO leads to the reduction of the ferric heme iron to the ferrous form and the breakage of the proximal iron-thiolate bond. According to Equation 1, the dimer-to-monomer conversion occurs prior to the reduction to the 5C ferrous form. This is consistent with prior reports of monomerization induced by the presence of NO, although we cannot exclude the less likely possibility of a direct reduction of the enzyme to the ferrous form prior to dissociation of the dimer. To determine whether NO causes any chemical modifications of the polypeptide chain of iNOSoxy, we carried out mass spectrometric measurements of the NO-treated samples. All samples examined were first subjected to trypsin digestion prior to mass spectrometric analysis. The major modification in the mass spectra of the NO-treated samples versus the control sample without NO treatment was the enhancement of the three fragment ions at m/z 581.94, 640.27, and 743.85 as shown in Fig. 5. The charge states of the three fragments were determined to be +3, +3, and +2, respectively, on the basis of their characteristic isotopic distributions. The parent masses of the ion peaks at m/z 581.94 and 743.85 (1742.82 and 1485.70 Da, respectively) calculated based on the charges are exact matches with two expected trypsin cleavage products of iNOSoxy corresponding to peptide fragments 82-97 and 393-404, respectively. These assignments were confirmed by the tandem mass spectrometric data (data not shown). Intriguingly, all observed fragment ion peaks in the mass spectra can be accounted for by the expected trypsin cleavage products, except the triply charged ion at m/z 640.27 with a parent mass of 1917.81 Da. We found that this ion peak is an exact match of peptide fragments 98-105 and 108-117 disulfide bond-linked through Cys104 and Cys109. This assignment was confirmed by the tandem mass data shown in supplemental Fig. S5. To further verify the disulfide-bonded peptide fragments, the trypsin-digested fragments of the NO-treated sample (in the presence of 4 m urea) were reduced by dithiothreitol (to reduce the disulfide bond) and alkylated by iodoacetamide (to alkylate the reduced free cysteine residues). This treatment resulted in the appearance of a doubly charged ion at m/z 553.76, the parent mass (1105.5 Da) of which is an exact match for peptide fragment 108-117 with a carbamido-methylated cysteine residue, at the expense of the fragment ion peak at m/z 640.27 (supplemental Fig. S6). The modified fragment 98-105 was not observed, possibly because of its low ionization propensity. These data further confirmed the presence of the disulfide-linked peptide fragment 98-105/108-117. It is important to note that, other than a very small contribution from a Cys109-Cys109 disulfide-bonded fragment (data not shown), no other disulfide-linked trypsin-digested iNOSoxy fragments were observed; furthermore, no fragments were found to contain any NO-derivatized amino acids. This is in contrast to the results of NO treatment of nitrophorins, in which the proximal cysteine bond becomes ruptured and nitrosylated (33Weichsel A. Maes E.M. Andersen J.F. Valenzuela J.G. Shokhireva T. Walker F.A. Montfort W.R. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 594-599Crossref PubMed Scopus (134) Google Scholar, 34Walker F.A. J. Inorg. Biochem. 2005; 99: 216-236Crossref PubMed Scopus (136) Google Scholar). Because the enzyme is in a monomeric state in the presence of 4 m urea, the formation of the disulfide-linked peptide fragment 98-105/108-117 must be a result of intramolecular rather than intermolecular interactions. Taken together, these data indicate that the NO-induced monomerization of the loose dimer is coupled to an intramolecular disulfide bond formation between cysteine residues at positions 104 and 109 and that the monomerization process exposes peptide fragments 82-97 and 393-404 to solvent, making them more accessible to trypsin digestion as reflected by the enhancement of the corresponding fragment ion peaks shown in Fig. 5. The data presented here clearly demonstrate that, in the absence of H4B, the ferric derivative of iNOSoxy is in equilibrium between a monomeric state and a loose dimeric state. Furthermore, l-Arg binding to the enzyme shifts the equilibrium toward the loose dimeric state, whereas the addition of H4B locks the enzyme in a tight dimeric state that resists NO-induced monomerization. The NO-induced monomerization in the loose dimer is associated with the formation of an intramolecular disulfide bond between Cys104 and Cys109, located in the zinc-binding motif in the dimer interface. The resulting monomeric enzyme with a 6C NO-bound heme iron gradually converts to a 5C NO-bound ferrous species because of the reduction of the heme iron and the concomitant breakage of the proximal iron-thiolate bond. Autoreduction Mechanism—One possible mechanism to account for the conversion of the 6C NO-bound ferric derivative to the five-coordinate NO-bound ferrous form is a heterolytic cleavage of the proximal iron-thiolate bond: Cys--Fe3+-NO→Cys·+Fe2+-NO. To test this mechanism, we re-examined the NO reaction with 4 m urea-treated iNOSoxy as a function of the NO concentration. We found that the formation rate of the 5C species increased approximately linearly as the NO concentration increased (data not shown). Because the Sn1-type heterolytic cleavage reaction predicts an NO concentration-independent kinetic process, this mechanism is excluded. NO-mediated conversion of a 6C NO-bound ferric protein to a 6C NO-bound ferrous protein has been well documented for histidine-ligated heme proteins such as hemoglobin and myoglobin (35Chien J.C. J. Am. Chem. Soc. 1969; 91: 2166-2168Crossref PubMed Scopus (89) Google Scholar, 36Addison A.W. Stephanos J.J. Biochemistry. 1986; 25: 4104-4113Crossref PubMed Scopus (99" @default.
• W1996652032 created "2016-06-24" @default.
• W1996652032 creator A5006206064 @default.
• W1996652032 creator A5050114481 @default.
• W1996652032 creator A5052285272 @default.
• W1996652032 creator A5059629802 @default.
• W1996652032 creator A5067430639 @default.
• W1996652032 creator A5071619668 @default.
• W1996652032 creator A5086969690 @default.
• W1996652032 date "2006-03-01" @default.
• W1996652032 modified "2023-09-29" @default.
• W1996652032 title "Regulation of the Monomer-Dimer Equilibrium in Inducible Nitric-oxide Synthase by Nitric Oxide" @default.
• W1996652032 cites W1479676079 @default.
• W1996652032 cites W1800678382 @default.
• W1996652032 cites W1970377569 @default.
• W1996652032 cites W1973018387 @default.
• W1996652032 cites W1973532480 @default.
• W1996652032 cites W1974109612 @default.
• W1996652032 cites W1976324923 @default.
• W1996652032 cites W1977533042 @default.
• W1996652032 cites W1988247358 @default.
• W1996652032 cites W1991861960 @default.
• W1996652032 cites W1992392409 @default.
• W1996652032 cites W2002109628 @default.
• W1996652032 cites W2004011343 @default.
• W1996652032 cites W2009402014 @default.
• W1996652032 cites W2013403540 @default.
• W1996652032 cites W2016668087 @default.
• W1996652032 cites W2017075380 @default.
• W1996652032 cites W2019743539 @default.
• W1996652032 cites W2019791075 @default.
• W1996652032 cites W2020736600 @default.
• W1996652032 cites W2021456770 @default.
• W1996652032 cites W2022669318 @default.
• W1996652032 cites W2026108937 @default.
• W1996652032 cites W2026623916 @default.
• W1996652032 cites W2033496106 @default.
• W1996652032 cites W2034091457 @default.
• W1996652032 cites W2035747601 @default.
• W1996652032 cites W2045639174 @default.
• W1996652032 cites W2045866331 @default.
• W1996652032 cites W2050094431 @default.
• W1996652032 cites W2050544473 @default.
• W1996652032 cites W2052105247 @default.
• W1996652032 cites W2055671467 @default.
• W1996652032 cites W2056420994 @default.
• W1996652032 cites W2057631848 @default.
• W1996652032 cites W2066311465 @default.
• W1996652032 cites W2067493659 @default.
• W1996652032 cites W2076291690 @default.
• W1996652032 cites W2076687382 @default.
• W1996652032 cites W2082876885 @default.
• W1996652032 cites W2084078405 @default.
• W1996652032 cites W2085742386 @default.
• W1996652032 cites W2091775196 @default.
• W1996652032 cites W2093735266 @default.
• W1996652032 cites W2104790006 @default.
• W1996652032 cites W2112413125 @default.
• W1996652032 cites W2113379061 @default.
• W1996652032 cites W2115549514 @default.
• W1996652032 cites W2121358285 @default.
• W1996652032 cites W2124306294 @default.
• W1996652032 cites W2124439270 @default.
• W1996652032 cites W2129097146 @default.
• W1996652032 cites W2145385387 @default.
• W1996652032 cites W2158917954 @default.
• W1996652032 cites W2160004875 @default.
• W1996652032 cites W2273844629 @default.
• W1996652032 cites W4234401654 @default.
• W1996652032 cites W83848988 @default.
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