Chimeras of Nitric-oxide Synthase Types I and III Establish Fundamental Correlates between Heme Reduction, Heme-NO Complex Formation, and Catalytic Activity
2001; Elsevier BV; Volume: 276; Issue: 26 Linguagem: Inglês
10.1074/jbc.m102509200
ISSN1083-351X
AutoresSubrata Adak, Kulwant S. Aulak, Dennis J. Stuehr,
Tópico(s)Eicosanoids and Hypertension Pharmacology
ResumoNeuronal nitric-oxide synthase (nNOS or NOS I) and endothelial NOS (eNOS or NOS III) differ widely in their reductase and nitric oxide (NO) synthesis activities, electron transfer rates, and propensities to form a heme-NO complex during catalysis. We generated chimeras by swapping eNOS and nNOS oxygenase domains to understand the basis for these differences and to identify structural elements that determine their catalytic behaviors. Swapping oxygenase domains did not alter domain-specific catalytic functions (cytochrome c reduction or H2O2-supportedN ω-hydroxy-l-arginine oxidation) but markedly affected steady-state NO synthesis and NADPH oxidation compared with native eNOS and nNOS. Stopped-flow analysis showed that reductase domains either maintained (nNOS) or slightly exceeded (eNOS) their native rates of heme reduction in each chimera. Heme reduction rates were found to correlate with the initial rates of NADPH oxidation and heme-NO complex formation, with the percentage of heme-NO complex attained during the steady state, and with NO synthesis activity. Oxygenase domain identity influenced these parameters to a lesser degree. We conclude: 1) Heme reduction rates in nNOS and eNOS are controlled primarily by their reductase domains and are almost independent of oxygenase domain identity. 2) Heme reduction rate is the dominant parameter controlling the kinetics and extent of heme-NO complex formation in both eNOS and nNOS, and thus it determines to what degree heme-NO complex formation influences their steady-state NO synthesis, whereas oxygenase domains provide minor but important influences. 3) General principles that relate heme reduction rate, heme-NO complex formation, and NO synthesis are not specific for nNOS but apply to eNOS as well. Neuronal nitric-oxide synthase (nNOS or NOS I) and endothelial NOS (eNOS or NOS III) differ widely in their reductase and nitric oxide (NO) synthesis activities, electron transfer rates, and propensities to form a heme-NO complex during catalysis. We generated chimeras by swapping eNOS and nNOS oxygenase domains to understand the basis for these differences and to identify structural elements that determine their catalytic behaviors. Swapping oxygenase domains did not alter domain-specific catalytic functions (cytochrome c reduction or H2O2-supportedN ω-hydroxy-l-arginine oxidation) but markedly affected steady-state NO synthesis and NADPH oxidation compared with native eNOS and nNOS. Stopped-flow analysis showed that reductase domains either maintained (nNOS) or slightly exceeded (eNOS) their native rates of heme reduction in each chimera. Heme reduction rates were found to correlate with the initial rates of NADPH oxidation and heme-NO complex formation, with the percentage of heme-NO complex attained during the steady state, and with NO synthesis activity. Oxygenase domain identity influenced these parameters to a lesser degree. We conclude: 1) Heme reduction rates in nNOS and eNOS are controlled primarily by their reductase domains and are almost independent of oxygenase domain identity. 2) Heme reduction rate is the dominant parameter controlling the kinetics and extent of heme-NO complex formation in both eNOS and nNOS, and thus it determines to what degree heme-NO complex formation influences their steady-state NO synthesis, whereas oxygenase domains provide minor but important influences. 3) General principles that relate heme reduction rate, heme-NO complex formation, and NO synthesis are not specific for nNOS but apply to eNOS as well. nitric oxide NO synthase calmodulin (6R)-5,6,7,8-tetrahydro-l-biopterin neuronal NOS endothelial NOS l-arginine flavin mononucleotide N ω-hydroxy-l-arginine a chimera containing an nNOS oxygenase domain, an eNOS reductase domain, and a CaM binding site a chimera containing an eNOS oxygenase domain, an nNOS reductase domain, and a CaM binding site polymerase chain reaction 4-(2-hydroxyethyl)-1-piperazinepropanesulfonic acid Nitric oxide (NO)1 is generated by nitric-oxide synthases (NOSs) and has multiple functions in physiology and pathology (1Craven S.E. Bredt D.S. Cell. 1998; 93: 495-498Abstract Full Text Full Text PDF PubMed Scopus (429) Google Scholar, 2MacMicking J. Xie Q.-W. Nathan C. Annu. Rev. Immunol. 1997; 15: 323-350Crossref PubMed Scopus (3484) Google Scholar, 3Dawson V.L. Dawson T.M. Prog. Brain. Res. 1998; 118: 215-229Crossref PubMed Google Scholar). Animals express three main NOS isoforms: one is cytokine-inducible and Ca2+-independent (iNOS or NOS II), and the two others are expressed constitutively (nNOS or NOS I; eNOS or NOS III) and become activated by Ca2+-dependent calmodulin (CaM) binding. All NOSs are bi-domain enzymes comprised of an N-terminal oxygenase domain that binds iron protoporphyrin IX (heme), (6R)-5,6,7,8-tetrahydro-l-biopterin (H4B), and l-arginine (Arg) and a C-terminal reductase domain that binds FMN, FAD, and NADPH (4Stuehr D.J. Biochim. Biophys. Acta. 1999; 1411: 217-230Crossref PubMed Scopus (813) Google Scholar, 5Roman L.J. Sheta E.A. Martasek P. Gross S.S. Lin Q. Masters B.S. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 8428-8432Crossref PubMed Scopus (244) Google Scholar, 6Hemmens B. Mayer B. Titheradge M.A. Methods in Molecular Biology. Humana Press, Totowa, NJ1997: 1-32Google Scholar). A CaM binding motif is located between the oxygenase and reductase domains, and its occupancy triggers electron transfer between the reductase domain FMN and the oxygenase domain heme (7Abu-Soud H.M. Stuehr D.J. Proc. Natl. Acad. Sci. U. S. A. 1993; 90: 10769-10772Crossref PubMed Scopus (398) Google Scholar). This enables NOSs to catalyze the NADPH- and O2-dependent oxidation of Arg to generate NO and citrulline, withN ω-hydroxyarginine (NOHA) being formed as an enzyme-bound intermediate (4Stuehr D.J. Biochim. Biophys. Acta. 1999; 1411: 217-230Crossref PubMed Scopus (813) Google Scholar, 5Roman L.J. Sheta E.A. Martasek P. Gross S.S. Lin Q. Masters B.S. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 8428-8432Crossref PubMed Scopus (244) Google Scholar, 6Hemmens B. Mayer B. Titheradge M.A. Methods in Molecular Biology. Humana Press, Totowa, NJ1997: 1-32Google Scholar). The constitutive NOSs share many features but differ markedly in their catalytic profiles. For example, nNOS is 3–4 times more active than eNOS in steady-state NO synthesis (8McCabe T.J. Fulton D. Roman L.J. Sessa W.C. J. Biol. Chem. 2000; 275: 6123-6128Abstract Full Text Full Text PDF PubMed Scopus (328) Google Scholar, 9Presta A. Liu J. Sessa W.C. Stuehr D.J. Nitric Oxide. 1997; 1: 174-187Crossref Scopus (53) Google Scholar, 10List B.M. Klosch B. Volker C. Gorren A.C. Sessa W.C. Werner E.R. Kukovetz W.R. Schmidt K. Mayer B. Biochem. J. 1997; 323: 159-165Crossref PubMed Scopus (140) Google Scholar, 11Martasek P. Liu Q. Liu J. Roman L.J. Gross S.S. Sessa W.C. Masters B.S. Biochem. Biophys. Res. Commun. 1996; 219: 359-365Crossref PubMed Scopus (142) Google Scholar, 12Nishida C.R. Ortiz de Montellano P.R. J. Biol. Chem. 1998; 273: 5566-5571Abstract Full Text Full Text PDF PubMed Scopus (95) Google Scholar, 13Abu-Soud H.M. Ichimori K. Presta A. Stuehr D.J. J. Biol. Chem. 2000; 275: 17349-17357Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). This holds true even though a majority of nNOS partitions into an inactive ferrous heme-NO complex immediately after initiating NO synthesis (14Abu-Soud H.M. Wang J. Rousseau D.L. Fukuto J. Ignarro L.J. Stuehr D.J. J. Biol. Chem. 1995; 270: 22997-23006Abstract Full Text Full Text PDF PubMed Scopus (201) Google Scholar). This lowers the concentration of active nNOS molecules by 4- or 5-fold, creates a condition in which oxidation of the ferrous-NO complex becomes rate-limiting, and shifts the apparentKm(O2) value of the enzyme to a much higher value (15Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 17434-17439Abstract Full Text Full Text PDF PubMed Scopus (54) Google Scholar, 16Abu-Soud H.M. Rousseau D.L. Stuehr D.J. J. Biol. Chem. 1996; 271: 32515-32518Abstract Full Text Full Text PDF PubMed Scopus (129) Google Scholar). In contrast, a very minor percentage of eNOS accumulates as a heme-NO complex during steady-state NO synthesis (13Abu-Soud H.M. Ichimori K. Presta A. Stuehr D.J. J. Biol. Chem. 2000; 275: 17349-17357Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar), and its slow catalysis is associated with a much slower heme reduction rate than nNOS (13Abu-Soud H.M. Ichimori K. Presta A. Stuehr D.J. J. Biol. Chem. 2000; 275: 17349-17357Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar, 17Roman L.J. Martasek P. Miller R.T. Harris D.E. de la Garza M.A. Shea T.M. Kim J.J.P. Masters B.S.S. J. Biol. Chem. 2000; 275: 29225-29232Abstract Full Text Full Text PDF PubMed Scopus (114) Google Scholar, 18Miller R.T. Martasek P. Omura T. Masters B.S.S. Biochem. Biophys. Res. Commun. 1999; 265: 184-188Crossref PubMed Scopus (74) Google Scholar). To better understand how the heme reduction rate controls NO complex formation and NO synthesis, we developed a kinetic simulation model for nNOS catalysis (19Santolini J. Adak S. Curran C.M.L. Stuehr D.J. J. Biol. Chem. 2001; 276: 1233-1243Abstract Full Text Full Text PDF PubMed Scopus (90) Google Scholar). Our model incorporates the key finding that newly synthesized NO binds to the ferric heme before it leaves the enzyme active site (20Boggs S. Huang L. Stuehr D.J. Biochemistry. 2000; 39: 2332-2339Crossref PubMed Scopus (66) Google Scholar). The kinetic model accurately simulates initial and steady-state features of nNOS catalysis including heme-NO complex formation, a concomitant deflection in NADPH oxidation and NO synthesis, and an increase in apparentKm(O2) value. Experimental evidence and additional simulations revealed that slowing down heme reduction in nNOS decreased the percentage of heme-NO complex and the rate of NO synthesis achieved in the steady state (21Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar). On this basis, we hypothesized that eNOS behavior might fit the kinetic model and suggested that the differences between nNOS and eNOS might be explained by their divergent heme reduction rates (19Santolini J. Adak S. Curran C.M.L. Stuehr D.J. J. Biol. Chem. 2001; 276: 1233-1243Abstract Full Text Full Text PDF PubMed Scopus (90) Google Scholar, 21Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar). To help test this hypothesis, we created chimeras by exchanging the oxygenase domains of nNOS and eNOS (NoxEred and EoxNred). In pioneering work, Ortiz de Montellano and co-workers (12Nishida C.R. Ortiz de Montellano P.R. J. Biol. Chem. 1998; 273: 5566-5571Abstract Full Text Full Text PDF PubMed Scopus (95) Google Scholar, 22Nishida C.R. Ortiz de Montellano P.R. J. Biol. Chem. 1999; 274: 14692-14698Abstract Full Text Full Text PDF PubMed Scopus (94) Google Scholar) generated similar chimeras from eNOS and nNOS and characterized their steady-state NO synthesis, cytochrome c reduction, and NADPH oxidation in response to Arg and H4B. The authors concluded that the reductase domain controlled the rates of NO synthesis and cytochrome c reduction, whereas oxygenase domains controlled NADPH oxidation in response to Arg and H4B. In our case, we hoped the chimeras would reveal how heme reduction, NO complex formation, and NO synthesis are related in eNOS and help us gauge to what extent reductase and oxygenase domains control these parameters in either NOS isoform. We examined flavin and heme reduction rates, heme-NO complex formation, and initial and steady-state catalytic behaviors of each chimera and compared these to data obtained with eNOS and nNOS. The results show how individual reductase and oxygenase domains regulate heme reduction and NO complex formation in eNOS and nNOS and how these two factors combine to regulate catalysis. All regents and materials were obtained from Sigma or sources reported previously (21Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar, 23Adak S. Ghosh S. Abu-Soud H.M. Stuehr D.J. J. Biol. Chem. 1999; 274: 22313-22320Abstract Full Text Full Text PDF PubMed Scopus (86) Google Scholar). Restriction digestions, cloning, bacterial growth, and the transformation and isolation of DNA fragments were performed using standard procedures. Rat nNOS DNA and bovine eNOS DNA were inserted into the 5′-NdeI and 3′-XbaI sites of the pCWori vector (23Adak S. Ghosh S. Abu-Soud H.M. Stuehr D.J. J. Biol. Chem. 1999; 274: 22313-22320Abstract Full Text Full Text PDF PubMed Scopus (86) Google Scholar, 24Ghosh S. Gachhui R. Crooks C. Wu C. Lisanti M.P. Stuehr D.J. J. Bio.l Chem. 1998; 273: 22267-22271Abstract Full Text Full Text PDF PubMed Scopus (135) Google Scholar). To create the chimeras we used site-directed mutagenesis to generate a unique restriction site between the end of the oxygenase domain and the beginning of the CaM binding domain in both bovine eNOS and rat nNOS. The unique restriction siteEco47III was incorporated at S485-A486 of eNOS and H714-V715 of nNOS. This created a silent mutation in eNOS and an His-Val→Ser-Ala mutation in nNOS. Sequence alignment using MacVector revealed that bothEco47III sites were located in identical positions in nNOS and eNOS. For making the Eco47III restriction site in eNOS, we used the QuikChangeTM site-directed mutagenesis kit from Stratagene. The oligonucleotides used to construct theEco47III site (underlined) in eNOS were synthesized by Integrated DNA Technologies, and their corresponding oligonucleotides were as follows: S485-A486-Eco47III sense, TGGAAAGGGAGCGCTACCAAGGGCGCCGGCATCA and S485-A486-Eco47III antisense, TGATGCCGGCGCCCTTGGTAGCGCTCCCTTTCCA. A RoboCycler gradient 96 from Stratagene was employed. The standard PCR cycling parameters were 3 min for denaturing of the template at 95 °C and 16 cycles for amplification (30 s for melting at 95 °C, 1 min for annealing at 60 °C, and 18 min for extension at 68 °C) followed by a 7-min extension at 68 °C. The protocol used ∼50 ng of template, 20 pmol of each primer, 2 μl of 10 mm dNTPs, and 1 μm 2.5-unit Pfu polymerase in a final volume of 100 μl. The PCR product was digested by 1 μl of DpnI endonuclease and then transformed into Epicurian Coli® XL1-Blue supercompetent cells. The Eco47III restriction site in the nNOS cDNA was constructed by subcloning a PCR-generated fragment from pCWori/nNOS using a 3′-oligo containing a newly engineeredEco47III site. The nNOS fragment was obtained by PCR amplification using Pfu Turbo DNA polymerase (Stratagene), which possesses higher fidelity than other polymerases. The nNOS cDNA fragment coding from the BlpI unique restriction site 622 to the SanDI restriction site 2162 was amplified using the following primers: primer 1, CCTGTGCTGAGCATCCTCAA; primer 2, TGGGGGTCCCGTTGGTGCCCTTCCAAGCGCTGGTGTTCCATGGATCAGG. Here the PCR cycling parameters were 3 min for denaturing of the template at 95 °C and 28 cycles for amplification (30 s for melting at 95 °C, 1 min for annealing at 58 °C, and 6 min for extension at 68 °C) followed by a 12-min extension at 68 °C. The protocol used ∼10 ng of template, 50 pmol of each primer, 2 μl of 10 mm dNTPs, and 1 μm 2.5-unit Pfupolymerase in a final volume of 100 μl. The PCR product and wild-type pCWori vector containing nNOS DNA were digested by both BlpI and SanDI restriction endonuclease enzymes, and fragments were isolated by 1% agarose gel. The double-digested fragment of wild-type NOS pCWori plasmid was replaced by the double-digested PCR fragment and transformed into JM109 cells to generate the recombinant plasmid. Both chimera proteins were constructed by interchanging the double restriction (NdeI and Eco47III)-digested fragments. Chimeric DNA constructs were confirmed by DNA sequencing at the Cleveland Clinic sequencing facility. Chimeric cDNAs in the pCWori plasmid were transformed into Escherichia coli strain BL21(DE3) for protein expression. Wild-type rat nNOS, bovine eNOS, and both chimera proteins (EoxNred and NoxEred) had a His6 tag attached to their N termini to aid purification. They were overexpressed inE. coli strain BL21(DE3) and purified by sequential chromatography on Ni2+-nitrilotriacetic acid and 2′,5′-ADP-Sepharose resins as described (15Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 17434-17439Abstract Full Text Full Text PDF PubMed Scopus (54) Google Scholar, 23Adak S. Ghosh S. Abu-Soud H.M. Stuehr D.J. J. Biol. Chem. 1999; 274: 22313-22320Abstract Full Text Full Text PDF PubMed Scopus (86) Google Scholar). The ferrous-CO adduct absorbing at 444 nm was used to quantitate heme protein content using an extinction coefficient of 74 mm−1cm−1 (A444–A500). Steady-state activities of wild-type and chimera proteins were determined separately at 25 °C using spectrophotometric assays that were described previously in detail (15Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 17434-17439Abstract Full Text Full Text PDF PubMed Scopus (54) Google Scholar, 23Adak S. Ghosh S. Abu-Soud H.M. Stuehr D.J. J. Biol. Chem. 1999; 274: 22313-22320Abstract Full Text Full Text PDF PubMed Scopus (86) Google Scholar). H2O2-dependent NOS oxidation of NOHA to nitrite was assayed in 96-well microplates at 25 °C as described previously (25Pufahl R.A. Wishnok J.S. Marletta M.A. Biochemistry. 1995; 34: 1930-1941Crossref PubMed Scopus (161) Google Scholar, 26Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 33554-33561Abstract Full Text Full Text PDF PubMed Scopus (214) Google Scholar) with modification. The assay volume was 100 μl and contained 40 mm EPPS, pH 7.6, 250 nm nNOS or mutant, 1 mm NOHA, 1 mmdithiothreitol, 25 units/ml superoxide dismutase, 0.5 mm EDTA, and 4 μm H4B. Reactions were initiated by adding 30 mm H2O2and stopped after 10 min by adding 1300 units of catalase. Nitrite was detected at 550 nm after adding the Griess reagent (100 μl) and quantitated based on nitrite standards. 1.0 μm NOS was diluted in an air-saturated 40 mmEPPS buffer, pH 7.6, containing 0.9 mm EDTA, 3 μm CaM, 200 μm dithiothreitol, 20 μm H4B, 160 μm NADPH, and 1 mm Arg in a final volume of 1 ml. Reactions were started by adding 1.2 mm Ca2+ and monitored by wavelength scanning at 15 °C. The kinetics of flavin and heme reduction were analyzed at 10 °C as described previously (15Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 17434-17439Abstract Full Text Full Text PDF PubMed Scopus (54) Google Scholar, 21Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar,26Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 33554-33561Abstract Full Text Full Text PDF PubMed Scopus (214) Google Scholar) using a diode array detector and stopped-flow apparatus from Hi-Tech Ltd. (model SF-61) equipped for anaerobic work. Heme reduction was followed by the formation of the ferrous-CO complex, and the kinetics were determined by an absorbance change at 444 nm. Reactions were initiated by rapidly mixing an anaerobic CO-saturated solution containing 50 μm NADPH with an anaerobic CO-saturated solution containing wild-type or mutant nNOS (2 μm), 40 mm EPPS buffer, pH 7.6, 10 μmH4B, 0.3 mm dithiothreitol, 5 mmArg, 4 μm CaM, and 1 mm Ca2+. Flavin reduction was monitored under the same conditions at 485 nm. Signal/noise ratios were improved by averaging at least 10 individual mixing experiments. The time course of absorbance change was fit to single or multiple exponential equations using a nonlinear least square method provided by the instrument manufacturer. Experiments were done at 10 °C using a diode array detector and stopped-flow apparatus from Hi-Tech Ltd. (model SF-61). To initiate NO synthesis, an air-saturated solution that contained 40 mm EPPS, pH 7.6, 2 μm nNOS or mutant, 0.4 mm dithiothreitol, 2 mm Arg, 20 μm H4B, 4 μm CaM or 20 μm soybean CaM proteins, 40 μm NADPH, and 0.5 mm EDTA was rapidly mixed with a buffered solution containing 2.4 mm Ca2+. Absorbance at 436 nm was monitored to follow ferrous heme-NO formation and absorbance at 340 nm was monitored to follow NADPH oxidation (15Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 17434-17439Abstract Full Text Full Text PDF PubMed Scopus (54) Google Scholar, 21Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google Scholar). The concentration of ferrous heme-NO complex formed during NO synthesis was estimated from the absorbance change at 436 nm using an extinction coefficient of 49,800 m−1 cm−1 (14Abu-Soud H.M. Wang J. Rousseau D.L. Fukuto J. Ignarro L.J. Stuehr D.J. J. Biol. Chem. 1995; 270: 22997-23006Abstract Full Text Full Text PDF PubMed Scopus (201) Google Scholar), and the amount of NADPH oxidation was determined using an extinction coefficient of 6,220 m−1 cm−1 at 340 nm. Signal/noise ratios were improved by averaging six consecutive scans. Each experiment was performed three separate times. Purified NoxEred and EoxNredboth exhibit the expected molecular mass (Fig.1). Spectroscopic analysis showed that their heme shifted to a high-spin state in the presence of 20 μm H4B and 1 mm Arg. Dithionite reduction of each chimera in the presence of Arg, H4B, and CO produced the expected 444-nm absorbance peak for the ferrous-CO complex in all cases (data not shown). These data confirm that exchanging the oxygenase domains of eNOS and nNOS did not alter protein expression or the physical properties of the oxygenase domains. We first compared reductase domain-independent catalysis by the chimeras and wild-type NOSs by measuring H2O2-dependent NOHA oxidation (Table I). The chimeras catalyzed NOHA oxidation to different degrees such that the activity of each chimera matched with the wild-type NOS that provided its oxygenase domain. This is consistent with the reaction not requiring electrons from the reductase domain (25Pufahl R.A. Wishnok J.S. Marletta M.A. Biochemistry. 1995; 34: 1930-1941Crossref PubMed Scopus (161) Google Scholar, 26Adak S. Wang Q. Stuehr D.J. J. Biol. Chem. 2000; 275: 33554-33561Abstract Full Text Full Text PDF PubMed Scopus (214) Google Scholar) and indicates that, under this circumstance, reductase domain identity did not influence catalysis by the oxygenase domains.Table IH2O2-dependent NOHA oxidation by wild-type NOSs and chimerasEnzymeNitrite/mole of enzymemolnNOS88 ± 2eNOS30 ± 2EoxNred26 ± 2NoxEred90 ± 5Incubations were run for 10 min at 25 °C prior to quenching as described under "Experimental Procedures." The values are the amount of product formed in 10 min and are the mean and S.D. for three measurements each. Open table in a new tab Incubations were run for 10 min at 25 °C prior to quenching as described under "Experimental Procedures." The values are the amount of product formed in 10 min and are the mean and S.D. for three measurements each. NADPH-dependent cytochromec reductase activity of each chimera in the presence or absence of CaM matched the activity of the NOS that provided its reductase domain (Table II). Thus, swapping oxygenase domains did not influence the reductase domain catalysis or response to CaM, which is consistent with previous results (12Nishida C.R. Ortiz de Montellano P.R. J. Biol. Chem. 1998; 273: 5566-5571Abstract Full Text Full Text PDF PubMed Scopus (95) Google Scholar, 22Nishida C.R. Ortiz de Montellano P.R. J. Biol. Chem. 1999; 274: 14692-14698Abstract Full Text Full Text PDF PubMed Scopus (94) Google Scholar). Steady-state NO synthesis activities of EoxNred and nNOS were identical and well coupled to NADPH oxidation (2.1 and 1.9 NADPH oxidized/NO formed, respectively) (Table II). In contrast, the steady-state NO synthesis activity of NoxEred was about one-third that of nNOS but was 33% greater than eNOS. NO synthesis by NoxEred and eNOS was also less coupled to their NADPH oxidation (4.4 and 4.0 NADPH oxidized/NO formed, respectively). Thus, the rates of NO synthesis and NADPH oxidation by each chimera equaled or approached the NOS isoform that provided its reductase domain.Table IICatalytic activities of wild-type NOSs and chimerasProteinCytochrome creductionNO synthesisNADPH oxidation+CaM−CaM+CaM−CaM+CaMmin−1min−1min−1Wild-type nNOS5600 ± 500452 ± 4057 ± 20110 ± 10Wild-type eNOS500 ± 3060 ± 215 ± 1060 ± 6EoxNred6000 ± 500500 ± 5056 ± 40120 ± 8NoxEred700 ± 10080 ± 1020 ± 2088 ± 8Activity measurements were performed at 25 °C in the absence or presence of CaM as described under "Experimental Procedures." Activities are expressed as moles of product formed/mole of heme/min. Values represent the mean and S.D. for three measurements. Open table in a new tab Activity measurements were performed at 25 °C in the absence or presence of CaM as described under "Experimental Procedures." Activities are expressed as moles of product formed/mole of heme/min. Values represent the mean and S.D. for three measurements. We measured the rates of NADPH-dependent flavin and heme reduction in CaM-bound chimeras using stopped-flow spectroscopy under anaerobic conditions. Fig. 2 (left panels) depicts flavin reduction as an absorbance decrease at 485 nm versustime. Flavin reduction was biphasic in both chimeras and was somewhat faster in the chimera containing the eNOS reductase domain (TableIII). This rate difference was also observed when comparing flavin reduction in eNOS and nNOS (13Abu-Soud H.M. Ichimori K. Presta A. Stuehr D.J. J. Biol. Chem. 2000; 275: 17349-17357Abstract Full Text Full Text PDF PubMed Scopus (115) Google Scholar). In contrast, heme reduction (as measured by CO binding) 2The initial absorbance decrease at 444 nm seen in the EoxNred trace is caused by flavin reduction (14Abu-Soud H.M. Wang J. Rousseau D.L. Fukuto J. Ignarro L.J. Stuehr D.J. J. Biol. Chem. 1995; 270: 22997-23006Abstract Full Text Full Text PDF PubMed Scopus (201) Google Scholar). in NoxEred was 380 times slower than in EoxNred (Fig. 2, right panels). The heme reduction rate observed for EoxNredclosely matched that reported for nNOS, and the rate seen with NoxEred was twice as fast as eNOS (TableIII).Table IIIObserved rate constants for NADPH-dependent flavin and heme reductionEnzymeFlavin reductionHeme reductionReferencek1k2s−1s−1eNOS85 ± 100.005 ± 0.00113Abu-Soud H.M. Ichimori K. Presta A. Stuehr D.J. J. Biol. Chem. 2000; 275: 17349-17357Abstract Full Text Full Text PDF PubMed Scopus (115) Google ScholarnNOS23 ± 2.13.4 ± 0.33.9 ± 0.321Adak S. Santolini J. Tikunova S. Wang Q. Johnson J.D. Stuehr D.J. J. Biol. Chem. 2001; 276: 1244-1252Abstract Full Text Full Text PDF PubMed Scopus (105) Google ScholarEoxNred22 ± 2.22.6 ± 0.33.8 ± 0.3In textNoxEred44 ± 3.22.6 ± 0.30.011 ± 0.001In textMeasurements were done with CaM-bound enzymes at 10 °C in a stopped-flow spectrophotometer under anaerobic conditions as described under "Experimental Procedures." Heme reduction was measured in the presence of CO. Rates are the averages obtained with two or three enzyme preparations. The data were best fit to a monophasic rate for heme reduction and a biphasic rate for flavin reduction. Open table in a new tab Measurements were done with CaM-bound enzymes at 10 °C in a stopped-flow spectrophotometer under anaerobic conditions as described under "Experimental Procedures." Heme reduction was measured in the presence of CO. Rates are the averages obtained with two or three enzyme preparations. The data were best fit to a monophasic rate for heme reduction and a biphasic rate for flavin reduction. We compared heme-NO complex buildup in the chimeras during steady-state NO synthesis. Fig.3 contains wavelength scans of nNOS, eNOS, and the two chimeras before and during NO synthesis at 15 °C. nNOS and EoxNred exhibited strong Soret absorbance positioned near 436 nm during steady-state NO synthesis, indicating their significant partitioning into a heme-NO complex. In contrast, NoxEred had a less prominent Soret absorbance at 436 nm in the steady state, indicating it formed less heme-NO complex, and eNOS showed very little absorbance gain in this region of the spectrum. Difference spectroscopy (Fig. 3,insets) confirmed that the heme-NO complex had a Soret peak at 436 nm and a broad visible absorbance at 560 nm in all cases, indicating that the heme-NO complex was predominantly ferrous. The estimated percentage of ferrous heme-NO complex present at steady state was ∼70% in nNOS and EoxNred, ∼25% in NoxEred, and ∼12% in eNOS. We next utilized stopped-flow spectroscopy to investigate the kinetics and extent of heme-NO complex formation and their relationship to NADPH oxidation during the initial and steady-state phases of NO synthesis. In Fig. 4, absorbance changes at 436 and 340 nm were monitored versus the time to follow heme-NO complex formation and NADPH oxidation, respectively, in reactions run at 10 °C. In all cases, heme-NO complex buildup was best described as a biphasic process (Table IV, k1, k2). The rates of heme-NO complex formation were essentially the same in nNOS and EoxNred, whereas they were somewhat faster in NoxEred compared with eNOS. The apparent k1 values for nNOS and EoxNred were four and six times faster than k1 values for NoxEred and eNOS, respectively. The apparent k2 values for nNOS and EoxNred were 14 and 8 times faster than k2 values for NoxEred and eNOS, respectively. The percentage of heme-NO complex at steady state was estimated from the stopped-flow data and di
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