NADPH oxidase 4 function as a hydrogen peroxide sensor

NADPH oxidase 4 function as a hydrogen peroxide sensor Abstract Nox4, a member of the NADPH- and oxygen-dependent oxidoreductases that generate reactive oxygen species (ROS), is widely expressed and constitutively active. To understand better its function and regulation, specific mutations in the Nox4 dehydrogenase (DH) domain were examined for effects on Nox4 oxidase activity. Transfection of His6-tagged Nox4 increased the amount of p22phox subunit in HEK293 cells, and a higher level of the heterodimer was observed in the nucleus-enriched fraction (NEF). NEF from Nox4-expressing HEK293 cells exhibited oxygen and H2O2 concentration-dependent NADPH oxidation rate. In Nox4-expressing cells, NEF and its partially purified form, the Nox4(P437H) mutant almost completely lost its oxidase activity, while Nox4(C546S), Nox4(C546L) or/and (C547L) had a significantly decreased rate of ROS production. The NADPH-dependent reduction of cytochrome c or cytochrome b5 by purified Nox4 DH domain was found regulated by the H2O2 concentration, and C546L and C547L mutants showed lower rates of the hemeprotein reduction. These conserved Cys residues in the DH domain respond to the cytosolic H2O2 concentration to regulate Nox4 activity. electron transfer, flavocytochrome, NADPH oxidase, reactive oxygen, redox regulation Nox enzymes comprise a family of NADPH oxidoreductases that generate reactive oxygen species (ROS) in a variety of cell types and tissues in response to numerous (both normal and abnormal) physiological signals (1, 2). The seven Nox isoforms (Nox1-5, Duox1 and Duox2) expressed in human tissues can be distinguished by their tissue distribution, structure and regulation, and reactive oxygen product. However, all seven Nox isoform proteins consist of highly conserved structural features as flavocytochrome; the C-terminal dehydrogenase (DH) domain contains binding sites for FAD and NADPH. The N-terminal transmembrane (TM) region consists of a six α-helical structure that involves four conserved histidine residues, located in the third and fifth TM helices that co-ordinate two heme groups (heme bL and bH). Regulated ROS production occurs at considerably reduced levels in most non-phagocytic tissues compared to activated phagocytes, or neutrophils (3, 4). Non-phagocytic Nox1 and Nox3 and phagocytic Nox2 form heterodimers with p22phox in membranes and require additional cytosolic regulatory proteins for function. Nox4 associates with p22phox in membranes but does not require cytosolic proteins. Nox5 and the dual oxidases Duox1 and Duox2 are activated by calcium (Ca2+) through a cytosolic EF-hand containing Ca2+ binding domain (5, 6). Nox1-3 and Nox5 generate primarily superoxide; Nox4, Duox1 and Duox2 produce mainly H2O2 (7–9). Nox4 may be unique among the Nox isoforms. Though Nox4 associates with p22phox in membranes, it does not require additional regulatory proteins or the proline-rich domain in p22phox, which is responsible for binding to the regulatory proteins, for activity (10, 11). Mutations in p22phox that inhibit Nox1-3 catalytic function do not alter Nox4-p22phox oxidase activity (12–14), suggesting that p22phox is required for Nox4 expression and specific membrane localization (15) but not for catalytic function. This has led to the concept that Nox4-dependent ROS generation is regulated primarily by its expression level (16–18). Moreover, most Nox/Duox isoforms are detected in many kinds of cells to produce ROS, and Nox4 is mainly observed in the membranes of endoplasmic reticulum, nucleus and mitochondria (19–22). In addition, unlike the other six isoforms, Nox4 is constitutively active (15, 23). Electron transfer activities of Nox1-5 DH domain proteins, Nox2 TM-Nox4 DH and Nox4 TM-Nox2 DH chimaera proteins were compared and only the Nox4 DH protein showed a significantly high rate without added activating subunits (8, 24), while Nox2 DH was activated by the association with p67phox-Rac1 complex (25). The DH domain of only Nox4 is constitutively ‘turned on’ compared with the DH domains of Nox1-3 and Nox5, thus, this property of the DH domain accounts for the constitutive active of the Nox4 holoenzyme (8). Here, to understand the mechanism of Nox4 regulation, mutations in the DH domain were examined for effects on oxidase activity. The present study suggested that H2O2-sensitive Nox4 DH domain containing cysteine residue(s) plays an important role in regulating ROS production in non-phagocytic cells. Experimental Procedures Materials Full-length cDNA encoding human Nox4 (amino acid residues 1-578) and N-terminally His6-tagged Nox4 cDNA were cloned into pcDNA3.1 and pCIG vectors, respectively (Invitrogen). cDNA encoding Nox-DH domains (Nox2 and Nox4) were subcloned into pMAL-C2X (New England Biolabs). DH domains corresponded to residues 283-570 for Nox2 and 304-578 for Nox4. These constructs, producing maltose binding protein (MBP)-tagged C-terminal Nox2 and Nox4 homologs, were expressed in Escherichia coli (E. coli) BL-21. The goat anti-rabbit IgG secondary anti-body linked to horseradish peroxidase (HRP) and pre-stained molecular weight markers for SDS-PAGE were from Bio-Rad. The polyclonal anti-body to a C-terminal peptide (residues 500–578) of Nox4 was from Novus Biologicals Inc., and monoclonal anti-bodies to MBP, amylose agarose and Factor Xa protease were from New England Biolabs. Mouse monoclonal anti-body 44.1 against human p22phox was from Santa Cruz Biotech, and mouse monoclonal anti-body #2366 to His6 was from Cell Signalling Technologies. The goat anti-mouse secondary anti-bodies linked to HRP came from Promega. Protease inhibitor cocktail (EDTA-free) and Amplex Red were from Roche and Invitrogen, respectively. Ferricytochrome c, glucose 6-phosphate DH (G6PDH), FAD, NADPH, NADP+, diphenylene iodonium (DPI), phenylmethanesulfonyl fluoride (PMSF), protein A-agarose Fast flow [50% (v/v)], 3, 3’-diaminobenzidine and nuclei EZ lysis buffer were purchased from Sigma Aldrich (St. Louis, MO). Site-directed mutagenesis and transient transfection of Nox4 According to the previously described methods (13), His6-tagged Nox4 point mutations Pro-437 mutated to His (denoted P437H), Cys-546 to Ser (C546S), Cys-546 to Leu (C546L) and Cys-547 to Leu (C547L) were generated using site-directed mutagenesis with human full length Nox4 cDNA. HEK293 cells were seeded at 1 x 106 cells/plate (10 cm diameter) and grown 24 h to 40–50% confluence in Dulbecco’s modified Eagle’s medium with 10% foetal bovine serum, 100 units/ml penicillin and 0.1 mg/ml streptomycin. Cells were transfected 48 h prior to use with mammalian expression vectors encoding His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4 (C546L/C547L) or empty vector, using FuGENE6 (Roche Molecular Biochemicals). Separation of nucleus-enriched fraction Transiently transfected HEK293 cells (3.5 ∼ 5 x 108 cells) cultured in several 10 cm tissue culture plates were harvested, washed twice with PBS, suspended in nuclei EZ lysis buffer, pH 7.4, containing 25 μM FAD and protease inhibitor cocktail (Complete Mini, Roche Diagnostics) plus 0.2 mM PMSF and disrupted using a glass homogenizer with a loose fitting pestle (3 min at 4°C). The homogenate was centrifuged at 800 x g, for 5 min in a KUBOTA 5922 rotor at 4°C to collect a nucleus-enriched fraction (NEF). The pooled NEF (30 μg) was subjected to 5–20% Tris-Glycine SDS-PAGE, electrotransferred to PVDF membranes (Millipore) and confirmed to be nucleus-enriched by immunoblotting with an anti-body to Lamin A. This fraction was also evaluated for oxygen or H2O2 concentration-dependent NADPH oxidase activity using fluorescence or absorption change. Western blot analysis His6-tagged Nox4 protein located in NEF of HEK293 cells was detected by Western blotting. Nucleus-enriched pellet extracts were prepared using the Nonidet P-40 lysis buffer containing 10% glycerol as described previously (9). Extracted proteins were separated on SDS-PAGE (5–20% gel) and transferred to an Immobilon PVDF membrane (Millipore). Proteins were visualized by incubation with primary anti-bodies overnight at 4°C with gentle shaking, and then with horseradish-linked secondary anti-body (1:3000 dilution, 2 h). Bands were detected by Luminescent Image Analyzer (Image Quant LAS 4000) after addition of Super Signal West Pico Chemiluminescent Substrate (Thermo Scientific), according to the manufacturer’s instructions. Purified MBP-tagged Nox2 DH, Nox4 DH, Nox4 DH (P437H), Nox4 DH (C546L) and Nox4 DH (C547L) proteins were subjected to 5–20% SDS-PAGE followed by immunoblotting using an anti-MBP anti-body and visualized as described above. Spectrophotometric measurement of Heme in NEF The NEF extracts were prepared using the Nonidet P-40 buffer according to the previously described method (9). The extracts prepared from HEK293 cells transfected with vector alone, vector encoding His6-Nox4, His6-Nox4(C546L), His6-Nox4 (C547L) or His6-Nox4(C546L/C547L) were used for the quantification of heme. Reduced minus oxidized difference spectra were recorded at 10 min intervals after addition of a few crystals of sodium dithionite until a stable spectrum was obtained. A molar absorption coefficient (ε426–408 nm) of 200 mM−1.cm−1 (26) for the Soret band was used for calculations. Superoxide generating activity Superoxide generation was assessed using superoxide dismutase (SOD)-inhibitable ferricytochrome c reduction quantified at 550 nm using an extinction coefficient of 21.1 mM−1.cm−1 (27). The indicated amount of HEK293 cell number, NEF or isolated Nox4 DH domain protein was added to the assay buffer [25 mM Hepes (pH 7.3) containing 25 μM FAD, 80 μM ferricytochrome c, 0.12 M NaCl, 3 mM KCl and 1 mM MgCl2], after which 50 μM NADPH was added and cytochrome c reduction was monitored in the presence and absence of 200 units of SOD at 25°C. Measurement of hydrogen peroxide generation Amplex Red (100 μM) plus 1.5 units/ml HRP was included in the assay mixture, and the reaction was initiated by the addition of intact cells, NEF or partially purified His6-Nox4 protein. The linear increase in the fluorescence of resorufin produced by oxidation of Amplex Red was measured (28). For experiments monitoring fluorescence, excitation and emission wavelengths of 572 nm and 583 nm, respectively, were used for microplate measurements. Reactions were monitored at 25°C for 10 min using Mithras LB940 multimode microplate reader (BERTHOLD Technologies). For NEF, 0.1 mg/ml final protein concentration was added to a reaction mixture containing 25 μM FAD, 50 μM glucose 6-phosphate, 50 μM NADP+ and commercial G6PDH (0.25 U/ml) was included. We used an NADPH-generating assay system consisting of glucose 6-phosphate, G6PDH and NADP+ to markedly decrease the background rate of Amplex Red oxidation by reduced pyridine nucleotide. The remaining low residual rate of enzyme-independent Amplex Red oxidation was then subtracted to obtain correct rates. The concentration of Amplex Red oxidized was calculated using an extinction coefficient of 54 mM−1•cm−1 at 572 nm (29), or when fluorescence was measured, using a standard curve generated from the addition of known amounts of hydrogen peroxide. Expression of Nox2 DH and Nox4 DH domains Truncated Nox DH clones were obtained by PCR using Nox cDNA cloned in the pMAL-C2X plasmid as the template. According to previously described methods (8, 24), PCR products for truncated Nox2 DH (residues 283-570), Nox4-DH (residues 304-578) and Nox4 DH mutant (residues 304-578, P437H, C546L or C547L) domains were purified using a PCR purification kit (Qiagen). The purified DNA fragments were ligated into the BamHI and HindIII restriction sites for the Nox2 DH domain, and BamHI and SalI for Nox4 DH domain in pMAL-C2X vector and transformed into E. coli. Transformants were selected from LB/ampicillin plates, and plasmids were isolated from 2 ml cultures as described previously (8). The plasmids were digested with restriction enzymes and separated on 1% agarose to confirm the presence of the insert. DNA sequences of the four clones were confirmed by nucleotide sequencing. Purification of expressed His6-Nox4 and MBP-Nox DH proteins His6-Nox4 or His6-Nox4 mutant was expressed in HEK293 cells and its partial purification was performed by a Ni-NTA affinity column chromatography as described previously (9). MBP-DH fusion proteins were induced in E. coli at 37°C by addition of 0.1 mM IPTG for 2.5 h and frozen at –80°C. Thawed cells were sonicated (3 x 10 s) and solubilized in 50 mM Hepes buffer, pH 7.5, containing 0.5 M NaCl, 1 mM PMSF, 1 mM EDTA, 1 mM dithiothreitol, protease inhibitor cocktail (1 μg/ml) and 0.15 M L-arginine at 3°C. Purification was performed by amylose-agarose column chromatography (10 x 15 mm) according to the methods described previously (8, 9). MBP-depletion was performed in the presence of Factor Xa for 6 h at 4°C according to the manufacturer’s instructions (New England Biolabs). MBP-fused and depleted proteins were used in 1 week to avoid gradual proteolysis and loss of activity. Purified MBP-fused and deleted Nox2-DH and Nox4 DH proteins were subjected to 5–20% SDS-PAGE followed by protein staining using Coomassie brilliant blue. Pyridine nucleotide-dependent electron transfer activity NADPH-dependent cytochrome c and cytochrome b5 reductase activities were assayed at 25°C according to the previously described methods (8). The activities were assayed in 1 ml volume of assay buffer [25 mM Hepes, pH 7.3, containing 0.12 M NaCl, 3 mM KCl, 1 mM MgCl2, 25 μM FAD, protease inhibitor cocktail (1 μg) and 80 μM electron acceptor]. After the mixture that included the purified Nox4 DH protein in the presence and absence of H2O2 had been pre-incubated for 5 min, the reaction was initiated by the addition of 50 μM NADPH. The reduction rates of electron acceptors were quantified by monitoring the absorbance changes at the appropriate wavelengths, and millimolar extinction coefficients for cytochrome c (21.1 mM−1•cm−1) (27) and cytochrome b5 (19 mM−1•cm−1) (30) at 556 nm were used to calculate the quantity of each electron acceptor reduced. Spectrophotometric measurements were performed using a Hitachi spectrophotometer with a temperature-controlled cuvette compartment. Oxygen concentration-dependent H2O2 generation by NEF The gas equilibration system consisted of a tightly capped 1.5 ml cuvette with a gas delivery needle and a gas exit needle that also served as a delivery port for addition of reagents. ROS measurement was conducted at 20°C in a total volume of 0.8 ml as described (9). The reaction mixture was equilibrated by gently bubbling (approximately one bubble per second) with the indicated percent of oxygen/nitrogen gas mixtures, with continuous gentle stirring for 10 min using a magnetic stirrer. Fluorescence increase at 583 nm due to Amplex Red oxidation was measured as described above. NADPH oxidation NADPH oxidation was measured spectrophotometrically at 340 nm in the presence of either NEF separated from HEK293 cells or purified MBP-Nox4 DH protein in 1 ml of the assay buffer as described above. After the assay mixture containing the sample was pre-incubated at 20°C for 5 min in the presence or absence of H2O2, the reaction was started by adding 50 μM NADPH. The amount of NADPH oxidized for 10 min was calculated using a molar absorption coefficient of 6.24 mM−1.cm−1 at 340 nm. The NADPH oxidation rate was also measured by the fluorescence decrease at 460 nm when excited at 340 nm. Results Analysis of C-terminal DH domain mutation of His6-Nox4 expressed in HEK293 cell To determine the importance of the NADPH DH domain in constitutive active Nox4 enzyme, point substitutions of proline residue at 437 and cysteine residue at 546 or/and 547 were generated by site-directed mutagenesis. Mutant His6-tagged Nox4 was expressed in HEK293 cells, and ROS-generating activity was monitored. Mutation of P437H resulted in nearly complete loss of Nox4 activity, whereas the C546S, C546L or/and C547L mutation indicated a remarkably decreased activity of wild-type Nox4 (Fig. 1A). Reactive oxygen product generation of His6-tagged Nox4 and mutant Nox4 (C546S, C546L or/and C547L) was verified to be largely inhibited by the general flavoprotein DH inhibitor, DPI. ROS generation was mainly detected in wild-type Nox4- and Nox4 mutant (C546S, C546L or/and C547L)-expressing cells, with the H2O2 identity validated due to inhibition by catalase and superoxide generating activity was greatly lower (Fig. 1B and C). The substitution of cysteine-546 to serine or leucine resulted in a significantly lower H2O2 and O2- producing activities as compared with wild-type Nox4 (Fig. 1B and C), and the mutation of either C547L or both C546L and C547L also showed a further decrease of ROS generating activity as observed in Fig. 1A–C, respectively. Consistent with His6-tagged Nox4, intact HEK293 cells stably expressing Nox4 also indicated that the major product was H2O2 and 40% or less superoxide was observed (9). These data also support that whole cells from His6-Nox4 transfected HEK293 cells were observed to generate mainly H2O2 at nearly the same rate as intact cells stably expressing native Nox4. Fig. 1 View largeDownload slide ROS generation by Nox4 in HEK293 cells—cDNA encoding His6-Nox4 (wild type), His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or empty vector alone (Mock) was transfected in HEK293 intact cells, as described in Experimental Procedures. (A) ROS was measured in the absence (white bars) and presence (grey bars) of 20 μM DPI using luminol chemiluminescence with 20 μM luminol and 0.32 unit HRP in 200 μl total volume. Results are shown as the mean ± SD. of three separate wells, and are representative of three separate transfection tests. (B) Hydrogen peroxide production was measured in intact His6-Nox4-expressing cells. The reaction was initiated by adding 5.2 x 105 cells to 0.8 ml of 25 mM Hepes, pH 7.3, containing 1.5 unit HRP and 100 μM Amplex Red. Amplex Red oxidation due to H2O2 was measured as the fluorescence increase at 583 nm when excited at 572 nm in the absence (white bars) and presence of 50 unit/ml of catalase (black bars). (C) Superoxide generation was quantified as superoxide dismutase (SOD)-inhibitable cytochrome c reduction at 550 nm, carried out in 1 ml of assay buffer, pH 7.3, containing 80 μM ferricytochrome c with or without SOD (final 300 U/ml) as described in Experimental Procedures. Values represent the mean activity ± SEM of three determinations using one set of transfected cells, and are representative of three transfection experiments. Fig. 1 View largeDownload slide ROS generation by Nox4 in HEK293 cells—cDNA encoding His6-Nox4 (wild type), His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or empty vector alone (Mock) was transfected in HEK293 intact cells, as described in Experimental Procedures. (A) ROS was measured in the absence (white bars) and presence (grey bars) of 20 μM DPI using luminol chemiluminescence with 20 μM luminol and 0.32 unit HRP in 200 μl total volume. Results are shown as the mean ± SD. of three separate wells, and are representative of three separate transfection tests. (B) Hydrogen peroxide production was measured in intact His6-Nox4-expressing cells. The reaction was initiated by adding 5.2 x 105 cells to 0.8 ml of 25 mM Hepes, pH 7.3, containing 1.5 unit HRP and 100 μM Amplex Red. Amplex Red oxidation due to H2O2 was measured as the fluorescence increase at 583 nm when excited at 572 nm in the absence (white bars) and presence of 50 unit/ml of catalase (black bars). (C) Superoxide generation was quantified as superoxide dismutase (SOD)-inhibitable cytochrome c reduction at 550 nm, carried out in 1 ml of assay buffer, pH 7.3, containing 80 μM ferricytochrome c with or without SOD (final 300 U/ml) as described in Experimental Procedures. Values represent the mean activity ± SEM of three determinations using one set of transfected cells, and are representative of three transfection experiments. The expression of Nox4 and its mutant proteins were examined by expressing His6-tagged Nox4 in HEK293 cells, followed by Western blotting (Fig. 2A). To determine if Nox4 and p22phox formed active heterodimers in NEF, we assessed the impact of p22phox expression on constitutively active Nox4. In NEF low levels of endogenous Nox4 and p22phox subunits are observed, but further expression of Nox4 subunit enhanced the steady-state level of p22phox. The results suggest that the transfection of Nox4 increased the amount of p22phox protein in HEK293 cells, and a higher level of the heterodimer was found in NEF. Almost similar levels of expression were detected for the four mutants compared with wild-type Nox4. Nox4 heterodimerizes with p22phox, and this association is assumed to be required for stability and location in the membrane of the nucleus. In accord with previous report (9), the increased Nox4 protein and ROS production were clearly confirmed in the NEF prepared from Nox4-expressing cells. Therefore, Nox4-enriched NEF was used in the present studies. Fig. 2 View largeDownload slide Determination of ROS level produced in a NEF or partially purified His6-Nox4 from HEK293 cells—(A) Expression of Nox4 and p22phox were determined in immunoblots of nuclear enriched fractions (NEF) (30 µg) prepared from HEK293 cells transfected with His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or Mock-transfected. (B) The NEF (0.1 mg) was supplemented with FAD, NADP+, glucose 6-phosphate and glucose 6-phosphate DH in NADPH-generating system, and hydrogen peroxide producing activity (white bars) was monitored by Amplex Red oxidation as described in Fig. 1B. Superoxide generation was assayed by SOD-inhibitable cytochrome c reduction monitored at 550 nm (grey bars). (C) NADPH-dependent H2O2 production of NEF was monitored in the absence (white bars) and presence (grey bars) of 20 μM DPI by measuring Amplex Red fluorescence change at 583 nm when excited at 572 nm. (D) Reduced minus oxidized difference spectra of NEF prepared from HEK293 cells. NEF expressed Mock (2.51 mg/ml), His6-Nox4(Wild) (2.05 mg/ml), His6-Nox4(C546L) (2.28 mg/ml) or His6-Nox4(C547L) (2.30 mg/ml) was reduced with a trace amount of Na2S2O4 for 5 min and difference spectra were measured at 20°C. (E) Partially purified His6-tagged forms (2.5 μg each) of Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L) were loaded onto SDS-PAGE gel (5–20%) and stained with Coomassie brilliant blue (top). NADPH-dependent H2O2 generating activity of purified His6-Nox4 was measured in the absence (white bars) and presence of either 20 μM DPI (grey bars) or 50 unit/ml of catalase (black bars) by monitoring Amplex Red fluorescence change at 583 nm as described above. Values represent the mean activity ± SEM of three determinations using one set of NEF (B, C) and purified protein (E). Similar results were obtained from three separate transfections for all experiments. Fig. 2 View largeDownload slide Determination of ROS level produced in a NEF or partially purified His6-Nox4 from HEK293 cells—(A) Expression of Nox4 and p22phox were determined in immunoblots of nuclear enriched fractions (NEF) (30 µg) prepared from HEK293 cells transfected with His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or Mock-transfected. (B) The NEF (0.1 mg) was supplemented with FAD, NADP+, glucose 6-phosphate and glucose 6-phosphate DH in NADPH-generating system, and hydrogen peroxide producing activity (white bars) was monitored by Amplex Red oxidation as described in Fig. 1B. Superoxide generation was assayed by SOD-inhibitable cytochrome c reduction monitored at 550 nm (grey bars). (C) NADPH-dependent H2O2 production of NEF was monitored in the absence (white bars) and presence (grey bars) of 20 μM DPI by measuring Amplex Red fluorescence change at 583 nm when excited at 572 nm. (D) Reduced minus oxidized difference spectra of NEF prepared from HEK293 cells. NEF expressed Mock (2.51 mg/ml), His6-Nox4(Wild) (2.05 mg/ml), His6-Nox4(C546L) (2.28 mg/ml) or His6-Nox4(C547L) (2.30 mg/ml) was reduced with a trace amount of Na2S2O4 for 5 min and difference spectra were measured at 20°C. (E) Partially purified His6-tagged forms (2.5 μg each) of Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L) were loaded onto SDS-PAGE gel (5–20%) and stained with Coomassie brilliant blue (top). NADPH-dependent H2O2 generating activity of purified His6-Nox4 was measured in the absence (white bars) and presence of either 20 μM DPI (grey bars) or 50 unit/ml of catalase (black bars) by monitoring Amplex Red fluorescence change at 583 nm as described above. Values represent the mean activity ± SEM of three determinations using one set of NEF (B, C) and purified protein (E). Similar results were obtained from three separate transfections for all experiments. In Fig. 2B, superoxide and hydrogen peroxide generation were compared in NEF from wild-type Nox4 and mutant Nox4-expressed cells. In the nuclear pellet fraction expressing Nox4, approximately 70% of ROS product was detected as H2O2, whereas 30% or less was observed as superoxide. Consistent with the whole cell assay as described in Fig. 1A and B, NEF expressing Nox4(P437H) almost completely lost H2O2 generation, and Nox4(C546L), Nox4(C547L) and Nox4(C546L/C547L) mutant released approximately 75%, 60% and 50% of H2O2, respectively, detected by wild-type Nox4 as the major ROS product. Inclusion of 20 μM DPI inhibited H2O2 generation more than 70% of wild-type Nox4 and Nox4(C546L, C547L or C546L/C547L) mutants (Fig. 2C). The concentration of heme in NEF prepared from non-transfected, His6-tagged Nox4, His6-Nox4(C546L), or His6-Nox4(C547L)-transfected HEK293 cells was 3.98, 10.9, 9.55 and 9.51 pmol/mg, respectively, indicating that the heme of wild-type His6-Nox4 and mutant Nox4(C546L or C547L) expressed in NEF was nearly the same level (Fig. 2D). In addition, the heme concentration in NEF from His6-Nox4(P437H)- or His6-Nox4(C546L/C547L)-transfected HEK293 was 10.5 and 9.63 pmol/mg protein, respectively. Therefore, the decrease in activity displayed by the Nox4 mutants was not due to altered protein expression, processing, or complex formation with p22phox. As shown in Fig. 2E, hydrogen peroxide generation of the partially purified His6-Nox4 was also compared in Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L). In accord with the results obtained by using NEF, purified Nox4(P437H) mutant almost completely lost H2O2 generating activity and the individual C546L and C547L mutations considerably decreased the oxidase activity. In addition, Nox4(C546L/C547L) double mutant showed a little higher inhibitory effect than either C546L or C547L mutant form (data not shown). These results suggest a possible critical role of proline 437, cysteine 546 and cysteine 547 residues in the Nox4 DH domain for H2O2 generation. Effect of oxygen or hydrogen peroxide on NADPH oxidase turnover in Nox4-expressed nucleus- enriched fraction of HEK293 cells In the previous study (9), we reported that Nox4 activity is regulated not only by its expression level, but also by oxygen availability, and that it therefore functions as an oxygen sensor. Nuclear membrane-associated Nox4 has a high Km value for oxygen that allows it to respond to physiological wide ranges of oxygen concentration, such that its enzymatic activity must be linked to an effect on signalling that can be translated into a cellular response. In fact, Nox2-dependent superoxide generation in intact human neutrophils shows a Km for oxygen of about 3%; on the other hand, Nox4 in intact cells indicated a higher oxygen Km value for H2O2 generation of around 15%, corresponding to the Km range seen for other known oxygen sensing enzymes (9). Here we found that Nox4(C546L or C547L) mutants have a higher Km value for oxygen of about 23%, as compared with a Km of 17% detected in Nox4-expressed NEF (Fig. 3A). Additionally, NADPH oxidation in Nox4-expressing NEF showed a higher rate compared to control NEF (Fig. 3B). The presence of 50 μM H2O2 significantly decreased the rate of NADPH oxidation in Nox4-expressed NEF, and the addition of 200 unit SOD also slightly reduced the oxidation rate. Consistent with these data, a decrease in the hydrogen peroxide concentration-dependent NADPH oxidation rate was detected in NEF prepared from Nox4 and Nox4-mutant (C546L or C547L) expressing HEK293 cells (Fig. 3C and D). NADPH oxidation was determined by measuring absorption and fluorescence changes of NADPH, which are thought to correspond to the H2O2 concentration-dependent effect on the rate of NADPH oxidation. In fact, the NADPH oxidation of NEF from wild-type Nox4-transfected cells was a little more remarkably decreased than mutant forms (C546L or C547L) in the presence of a lower range of H2O2 concentrations (0–100 μM). In the presence of 50 μM H2O2, NADPH oxidase activity of NEF expressed wild-type Nox4 was approximately 52% decrease, while Nox4 (C546L) or (C547L) mutant indicated almost same decrease, 45 or 44%, respectively. Thus, the catalytic function of NADPH-dependent Nox4 enzyme appears to be closely regulated within the physiological range of cytosolic H2O2 concentrations as well as cellular pO2 values. Fig. 3 View largeDownload slide Hydrogen peroxide and oxygen concentration-dependent NADPH oxidation activity of NEF expressing His6-Nox4—(A) Oxygen concentration-dependent H2O2 production by NEF stably expressing His6-Nox4 (white circles), His6-Nox4(C546L) (filled circles) or His6-Nox4(C547L) (white triangles) was measured using Amplex Red fluorescence at 583 nm. A double-reciprocal plot of initial velocity versus O2 concentration on the top was formed to calculate Km and Vmax values. (B) After the NEF was preincubated for 5 min at 20°C in the presence of either H2O2 or SOD, 50 μM NADPH was added. The NADPH oxidation rate was monitored by measuring its absorption decrease at 340 nm without (white bars) or with added either 50 μM H2O2 (grey bars) or 200 U/ml SOD (filled bars). The values shown in (A) and (B) are the mean ± SEM of three determinations from one set of transfections, and is representative of three experiments repeated using three different sets of transfected cells. (C) NEF (78.5 μg) prepared from His6-Nox4- (white circles), His6-Nox4(C546L)- (filled circles) or His6-Nox4(C547L)-expressed cells (white triangles) was suspended in 0.8 ml of 25 mM Hepes, pH 7.3, containing 1 mM MgCl2 and the indicated concentrations of H2O2. After the NEF suspension was preincubated for 5 min at 20°C, 50 μM NADPH was added and measured the absorption change at 340 nm. (D) The rate of NADPH oxidation was monitored by fluorescence change at 460 nm when excited at 340 nm. The each symbol indicates the same NEF prepared from HEK293 as described in Fig. 3C. Data points and error bars in (C) and (D) indicate the mean ± SEM of three determinations from single assays, and the experiments shown are representative of three different experiments. Fig. 3 View largeDownload slide Hydrogen peroxide and oxygen concentration-dependent NADPH oxidation activity of NEF expressing His6-Nox4—(A) Oxygen concentration-dependent H2O2 production by NEF stably expressing His6-Nox4 (white circles), His6-Nox4(C546L) (filled circles) or His6-Nox4(C547L) (white triangles) was measured using Amplex Red fluorescence at 583 nm. A double-reciprocal plot of initial velocity versus O2 concentration on the top was formed to calculate Km and Vmax values. (B) After the NEF was preincubated for 5 min at 20°C in the presence of either H2O2 or SOD, 50 μM NADPH was added. The NADPH oxidation rate was monitored by measuring its absorption decrease at 340 nm without (white bars) or with added either 50 μM H2O2 (grey bars) or 200 U/ml SOD (filled bars). The values shown in (A) and (B) are the mean ± SEM of three determinations from one set of transfections, and is representative of three experiments repeated using three different sets of transfected cells. (C) NEF (78.5 μg) prepared from His6-Nox4- (white circles), His6-Nox4(C546L)- (filled circles) or His6-Nox4(C547L)-expressed cells (white triangles) was suspended in 0.8 ml of 25 mM Hepes, pH 7.3, containing 1 mM MgCl2 and the indicated concentrations of H2O2. After the NEF suspension was preincubated for 5 min at 20°C, 50 μM NADPH was added and measured the absorption change at 340 nm. (D) The rate of NADPH oxidation was monitored by fluorescence change at 460 nm when excited at 340 nm. The each symbol indicates the same NEF prepared from HEK293 as described in Fig. 3C. Data points and error bars in (C) and (D) indicate the mean ± SEM of three determinations from single assays, and the experiments shown are representative of three different experiments. Hydrogen peroxide concentration-dependent electron transferase activity of constitutive active Nox4 DH domain Herein, we explore the hypothesis that the structural features necessary for constitutively active Nox4 reside in a cytosol-facing DH domain that contains a binding site for NADPH and one for FAD. To determine whether this domain was sufficient to exhibit spontaneous electron transferase, the Nox4 DH domain was expressed and purified as an MBP fusion protein and electron transfer activity toward cytochrome c or cytochrome b5 was measured (Fig. 4). In addition, to investigate the structural features necessary for constitutive activity are ascribed to the Nox4 DH domain, Nox2 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domains were also expressed and purified to compare their NADPH-dependent electron transfer activities. Purified MBP-Nox DH domain fusion and MBP-depleted Nox DH proteins corresponded in size to their predicted molecular masses on SDS-PAGE (Fig. 4A-a, -b) and MBP-fused Nox DH proteins were recognized on Western blots using an anti-MBP anti-body (Fig. 4A–c). As previously reported (8), the MBP-Nox4 DH domain fusion protein showed significant NADPH-dependent electron transfer activity toward one-electron acceptor heme proteins (Fig. 4B). The MBP-Nox4 DH(C546L or C547L) mutant proteins showed a distinctive decrease in NADPH-dependent electron transferase activity, and the turnover number toward each electron acceptor was about 50–60% level observed for the wild-type MBP-Nox4 DH. In addition, Nox2 DH exhibited a negligible NADPH-dependent electron transferase activity, and the electron transfer rate of Nox4 DH(P437H) domain was very low for each electron acceptor (<10 min−1). As shown in Fig. 4C, MBP-depleted Nox2 DH and Nox4 DH(P437H) showed little or no electron transfer activity, similar to the results observed for the MBP-fused forms. Although MBP-depleted Nox4 DH and Nox4 DH mutant (C546L or C547L) catalyzed the NADPH-dependent reduction of cytochrome c or cytochrome b5, respectively, their turnover rates were a little lower than the values observed for MBP-fused Nox4 DH, Nox4 DH(C546L) and Nox4(C547L) proteins. MBP alone showed no activity and the MBP-fused forms of the Nox4 DH domains were used in the present study because of the poor solubility and lower yield of the DH domains when the MBP tag was cleaved and deleted. Little or no Nox4 DH domain-dependent electron transferase activity was observed in the absence of added FAD, indicating loss of FAD during the purification. Fig. 4 View largeDownload slide NADPH-dependent electron transfer activities of wild and mutant MBP-fused Nox DH domain expressed in E. coli—(A) Purified MBP-fusion forms of Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domain (1 μg protein each) were loaded onto 5–20% (w/v) SDS-PAGE gel and stained with Coomassie brilliant blue (a). MBP-depleted form of purified DH proteins (1.5 μg each) were subjected to 5–20% (w/v) SDS-PAGE, followed by protein stain (b). Purified MBP-Nox DH proteins were immunoblotted with anti-body to MBP (c). (B) Pyridine nucreotide-dependent electron transferase activities of purified MBP-fused Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L), Nox4 DH(C547L) and MBP alone were assayed for cytochrome c (white bars) and cytochrome b5 (grey bars) reductase activities. Each reductive activity indicates the average of three independent assays, with the error bars showing the SD. (C) NADPH-dependent electron transfer activities of MBP-depleted Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. The electron transfer assay condition, electron acceptors used and each activity presentation were the same as described in Fig. 4B. Fig. 4 View largeDownload slide NADPH-dependent electron transfer activities of wild and mutant MBP-fused Nox DH domain expressed in E. coli—(A) Purified MBP-fusion forms of Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domain (1 μg protein each) were loaded onto 5–20% (w/v) SDS-PAGE gel and stained with Coomassie brilliant blue (a). MBP-depleted form of purified DH proteins (1.5 μg each) were subjected to 5–20% (w/v) SDS-PAGE, followed by protein stain (b). Purified MBP-Nox DH proteins were immunoblotted with anti-body to MBP (c). (B) Pyridine nucreotide-dependent electron transferase activities of purified MBP-fused Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L), Nox4 DH(C547L) and MBP alone were assayed for cytochrome c (white bars) and cytochrome b5 (grey bars) reductase activities. Each reductive activity indicates the average of three independent assays, with the error bars showing the SD. (C) NADPH-dependent electron transfer activities of MBP-depleted Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. The electron transfer assay condition, electron acceptors used and each activity presentation were the same as described in Fig. 4B. Consistent with previous studies (8, 24), Nox4 DH domain-dependent activities were very low using NADH rather than NADPH as an electron donor. NADPH-dependent cytochrome c or cytochrome b5 reduction by purified Nox4 DH was not largely affected by added SOD (Fig. 5A and B), and is therefore due to a direct electron transfer from the enzyme-bound FAD rather than a superoxide-mediated reaction. The turnover rate of MBP-Nox4 DH(C546L or C547L) also was not remarkably decreased in the presence of added SOD. Addition of DPI and H2O2 strongly decreased NADPH-specific hemeprotein reductase activities of the point mutant Nox4 DH(C546L or C547L) domain as well as wild-type Nox4 DH. In addition, the NADPH concentration-dependent cytochrome c reduction rates of MBP-Nox4 DH and MBP-Nox4 DH mutant (C546L or C547L) were measured (Fig. 5C). The Km value for NADPH was determined to be approximately 17 ± 2 μM in both wild and mutant DH proteins, a value close that previously reported for Nox4 DH (20 ± 5 μM) (8). On the other hand, Vmax values for Nox4 DH(C546L) and Nox4 DH(C547L) mutants were markedly decreased and determined to be 80 ± 5 and 65 ± 5 nmol cytochrome c reduced/min/mg protein, respectively, as compared with Vmax (200 ± 10) of wild-type Nox4 DH. The addition of 50 μM H2O2 considerably decreased the NADPH oxidation rates of MBP-fused Nox4 DH and Nox4 DH mutant (C546L or C547L) proteins in the presence of cytochrome c, whereas the inhibitory effect of H2O2 was significantly lost in the presence of catalase (Fig. 5D). In addition, it was noticed that NADPH oxidative activity of purified Nox4 DH(C547L) mutant was further decreased, about 26% lower than Nox4 DH(C546L) in the absence and presence of 50 μM H2O2 (Fig. 5D). The NADPH oxidation of MBP-Nox4 DH linked with cytochrome b5 reduction was also decreased in the presence of H2O2 (data not shown). Thus, the Nox4 DH domain includes a H2O2-sensing region, and NADPH-specific cytochrome c or cytochrome b5 reductase activity of this domain is clearly regulated by the local H2O2 concentration. Fig. 5 View largeDownload slide Effects of SOD, DPI and H2O2 on NADPH-dependent electron transfer activity by MBP-Nox4 DH or MBP-Nox4 DH mutant—(A) Cytochrome c reduction at 550 nm was measured as described in Figure 4B in the absence and presence of 300 U/ml SOD, 20 μM DPI or 50 μM H2O2. Purified MBP-Nox4 DH (white bars), MBP-Nox4 DH(C546L) (grey bars) or MBP-Nox4 DH(C547L) (black bars) protein was subjected to the assay as indicated. (B) The reduction of cytochrome b5 was quantified by monitoring the absorbance change at 556 nm without and with added SOD, DPI or H2O2 at the indicated concentrations in Fig. 5A. (C) NADPH concentration-dependent cytochrome c reduction of purified MBP-Nox4 DH or MBP-Nox4 DH mutants. Nox4 DH (○), Nox4 DH(C546L, •) and Nox4 DH(C547L, ▵)were assayed for cytochrome c reductase activity, and a double-reciprocal plot of initial velocity versus NADPH concentration on the top was used to determine Km and Vmax values. (D) After purified MBP-Nox4 DH or its mutant form was preincubated with 25 μM FAD and 80 μM cytochrome c in the absence (white bars) or presence of 50 μM H2O2 (grey bars) for 5 min, 50 μM NADPH was added and the rate of NADPH oxidation at 340 nm was assayed at 25°C. NADPH oxidation was also measured in the presence of 50 μM H2O2 just after adding 130 unit/ml of catalase (black bars). (A–D) The data are representative of three separate experiments, with the error bars showing the SD. Fig. 5 View largeDownload slide Effects of SOD, DPI and H2O2 on NADPH-dependent electron transfer activity by MBP-Nox4 DH or MBP-Nox4 DH mutant—(A) Cytochrome c reduction at 550 nm was measured as described in Figure 4B in the absence and presence of 300 U/ml SOD, 20 μM DPI or 50 μM H2O2. Purified MBP-Nox4 DH (white bars), MBP-Nox4 DH(C546L) (grey bars) or MBP-Nox4 DH(C547L) (black bars) protein was subjected to the assay as indicated. (B) The reduction of cytochrome b5 was quantified by monitoring the absorbance change at 556 nm without and with added SOD, DPI or H2O2 at the indicated concentrations in Fig. 5A. (C) NADPH concentration-dependent cytochrome c reduction of purified MBP-Nox4 DH or MBP-Nox4 DH mutants. Nox4 DH (○), Nox4 DH(C546L, •) and Nox4 DH(C547L, ▵)were assayed for cytochrome c reductase activity, and a double-reciprocal plot of initial velocity versus NADPH concentration on the top was used to determine Km and Vmax values. (D) After purified MBP-Nox4 DH or its mutant form was preincubated with 25 μM FAD and 80 μM cytochrome c in the absence (white bars) or presence of 50 μM H2O2 (grey bars) for 5 min, 50 μM NADPH was added and the rate of NADPH oxidation at 340 nm was assayed at 25°C. NADPH oxidation was also measured in the presence of 50 μM H2O2 just after adding 130 unit/ml of catalase (black bars). (A–D) The data are representative of three separate experiments, with the error bars showing the SD. Discussion Purified Nox4 DH-dependent reduction of heme protein such as cytochrome c or cytochrome b5 supports a model for the endogeneous NADPH-to-FAD-to-heme bL-to-heme bH electron transfer that occurs in the holoenzyme of Nox4 flavocytochrome. In this model, we were able to indicate the correspondence in H2O2 effect on intact Nox4 turnover during the NADPH-dependent ROS formation (Fig. 6A) and NADPH-dependent electron transfer rates to artificial hemeproteins (Fig. 6B), suggesting that cytosolic H2O2 regulates the rate-limiting FAD reduction in holo-Nox4 enzyme. Thus, the present data provide a reaction system in which H2O2 concentration-dependent reversible change of Nox4 DH domain structure appears to be co-regulated by peroxidases such as TRx, GRx and catalase (Fig. 6C). Fig. 6 View largeDownload slide Hydrogen peroxide-dependent regulation of constitutive active Nox4 enzyme—(A) Hydride ion transfer from NADPH to FAD is assumed to be the rate-determining step in Nox4 flavocytochrome, which is regulated by cytosolic H2O2 concentration. After the reduction of FAD by NADPH, the DH domain/TM-heme domain interaction may help orient and localize the FAD in close proximity to heme bL, thus allowing rapid electron transfer. (B) The electron transfer rate from purified Nox4 DH toward artificial heme proteins is affected in the presence of physiological range of H2O2. (C) Nox4 activity is affected by H2O2 concentration-dependent Cys-SH residue(s) oxidation of DH domain, which is adjusted by cytosolic peroxidases such as thioredoxin (TRx), glutaredoxin (GRx), or catalase. Fig. 6 View largeDownload slide Hydrogen peroxide-dependent regulation of constitutive active Nox4 enzyme—(A) Hydride ion transfer from NADPH to FAD is assumed to be the rate-determining step in Nox4 flavocytochrome, which is regulated by cytosolic H2O2 concentration. After the reduction of FAD by NADPH, the DH domain/TM-heme domain interaction may help orient and localize the FAD in close proximity to heme bL, thus allowing rapid electron transfer. (B) The electron transfer rate from purified Nox4 DH toward artificial heme proteins is affected in the presence of physiological range of H2O2. (C) Nox4 activity is affected by H2O2 concentration-dependent Cys-SH residue(s) oxidation of DH domain, which is adjusted by cytosolic peroxidases such as thioredoxin (TRx), glutaredoxin (GRx), or catalase. Although Nox4 expression is greatest in the kidney (23, 31, 32), it is widely expressed in many other cell types (32–34) and hence may have a cellular function that is more general than that of other Nox enzymes with more restricted tissue expression. We found that transfection of His6-tagged Nox4 cDNA to HEK293 cells induced further synthesis of p22phox subunit and increased Nox4-p22phox complex was observed in nuclear membrane fraction (Fig. 2A), which agrees with previous reports that stabilized p22phox expression responds to the level of constitutively active Nox4 formation and its intracellular localization (35). Thus, the Nox4 flavocytochrome-p22phox interaction produces a more stable conformation of the Nox4 protein localized to the nuclear membranes of HEK293 cells for the catalytic function to produce ROS. A recent study reported the major product from Nox4 is H2O2, and its activity is regulated by both its expression level and oxygen availability, functioning as an oxygen sensor (9). In fact, Nox4 generated H2O2 approximately in direct proportion to oxygen at concentrations below about 10%, making it a sensitive reporter of tissue oxygenation level. That Nox4 shows a high Km value for oxygen and generates mostly hydrogen peroxide indicates that it responds to the output of H2O2 rather than to external signals via intermediate signalling mechanisms, such as changes in cellular Ca2+ concentration or phosphorylation of regulatory subunits. Hydrogen peroxide shows relative specificity toward low pKa cysteine residues allowing its use as a signalling molecule to regulate enzyme activity (36, 37). We suggest that H2O2 participates in signalling ROS to regulate Nox4 activity in adaptive response to oxidative stress or hypoxic condition. Several studies indicate that specific location of Cys residues in proteins is associated with subcellular function (38, 39), suggesting a basic consideration of framework for differential protein oxidation due to oxidative stresss and toxicity linked with redox response and function. The reactive thiol side chain of Cys residues can function as a sensor or switch, changing between the reduced and oxidized state in response to fluctuations in ROS. Depending on the local ROS concentration, hydrogen peroxide, superoxide or NO can react with Cys residue to form reversible modifications (S-sulfenylation, disulfide bond formation, or S-nitrosylation). The introduction of new disulfide formation has the potential significantly to alter protein conformation and affect bio-function. Many proteins with reactive cysteines are involved in controlling thiol-disulfide exchange reactions to regulate enzyme activity and maintain cellular redox balance (40–42). In the steady state, thiols undergo oxidation due to increased oxidant such as H2O2 produced by Nox enzymes, suggesting that reactive peptidyl cysteine oxidation is associated with functional signal networks (43). Conceivably, in the nuclei, reversible structural change of redox-sensitive protein has a critical role for the activation or inactivation of transcriptional factors (44, 45). Hydrogen peroxide can cross membranes and is relatively stable. Although hydrogen peroxide has been intrinsically associated with oxidative stress, the physiological role of this mild oxidant may be as a redox signal messenger, and its production is likely controlled by the H2O2-sensitive Nox4 enzyme (46, 47). In fact, the redox sensitive thiol side chain oxidation of Cys residues in Nox4 DH domain appears to affect the Nox4 activity. Since NADPH-dependent electron transfer rate of the DH domain to heme protein such as cytochrome c or cytochrome b5 was remarkably affected by H2O2, the reversible oxidative modification of conserved and reactive Cys residues by H2O2 are assumed to be responsive to Nox4 activity. The NADPH oxidation rate of either wild-type or mutant Nox4 expressed in the membranes of NEF was clearly decreased with increasing H2O2 concentration (Fig. 3C and D). On the other hand, NADPH-dependent reduction of the artificial electron acceptor by either Nox4(C546L) or Nox4(C547L) mutant DH was significantly lower rate and less sensitive to H2O2 (Fig. 5A and B). In accord with these results of the mutant Nox4 DH, the decrease of H2O2-producing activity linked with NADPH oxidation rate of Nox4(C546L or C547L) flavocytochrome was also implied in NEF (Figs 2C and 3B). Our results, therefore, suggest that H2O2 production rate by constitutively active Nox4 should be controlled in intracellular H2O2 concentration. The increased H2O2 production with time in the nuclei caused by Nox4 up-regulation may induce oxidative nuclear damage leading to dysfunction and apoptotic cell death; therefore, the activity of Nox4 should be constantly auto-regulated by its DH flavoprotein domain. Since H2O2 reacts with Fe2+ and produces hydroxy radical (.OH) which enhances the risk of intracellular damages, it is important to regulate the H2O2 generating activity of Nox4 enzyme. On the other hand, anti-oxidant enzymes such as catalase, thioredoxin and glutaredoxin are considered to preferentially reduce sulfenic acid or disulfide bridge formation of Cys residues in DH domain of Nox4 to keep thermodynamic stability of the enzyme activity (Fig. 6C). In view of the three-dimensional protein model on the basis of its primary structure (48), the Nox4 C-terminal DH domain was shown to be composed of two units, a β-fold capped by a single helix to bind FAD, followed by a Rossmann-fold that binds NADPH (Fig. 7). In this model, Cys546 or/and Cys547 are suggested to be part of the NADPH binding site based on homology to ferredoxin-NADP+ reductase, cytochrome P-450 reductase and nitric oxide synthase (49, 50). When the C-terminal conserved amino acid sequences of NADPH binding sites are compared among human Nox family members (34), the Cys546-Cys547 sequence is located specifically in only the Nox4 DH domain (Table I). Thus, the hydrogen peroxide-dependent sulfenic acid or disulfide formation of these Cys residues may induce a limited reversible change of the DH domain conformation affecting the ROS producing turnover of constitutively active Nox4 enzyme. Table I. C-terminal conserved putative NADPH-binding amino acid sequences of human Nox isoform— Underline indicates Nox4 DH specific conserved Cys546/Cys547 residues that are predicted to control NADPH DH activity in response to intracellular H2O2 concentration Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Table I. C-terminal conserved putative NADPH-binding amino acid sequences of human Nox isoform— Underline indicates Nox4 DH specific conserved Cys546/Cys547 residues that are predicted to control NADPH DH activity in response to intracellular H2O2 concentration Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Fig. 7 View largeDownload slide Three-dimensional model of Nox4—A three-dimensional model is given for Nox4 based on its primary structure (48). The Nox4 C-terminal DH domain is composed of two units, a β-fold capped by a single helix that binds FAD, and a Rossman-fold that binds NADPH. Cys546 and Cys547 are localized in NADPH binding site. Presumably, the hydrogen peroxide-dependent sulfenic acid or disulfide formation between these two Cys residues may induce a limited reversible change in the conformation of the DH domain that affects turnover of the ROS-producing Nox4 enzyme. Fig. 7 View largeDownload slide Three-dimensional model of Nox4—A three-dimensional model is given for Nox4 based on its primary structure (48). The Nox4 C-terminal DH domain is composed of two units, a β-fold capped by a single helix that binds FAD, and a Rossman-fold that binds NADPH. Cys546 and Cys547 are localized in NADPH binding site. Presumably, the hydrogen peroxide-dependent sulfenic acid or disulfide formation between these two Cys residues may induce a limited reversible change in the conformation of the DH domain that affects turnover of the ROS-producing Nox4 enzyme. Acknowledgement The authors express appreciate to Dr Toshitsugu Yubisui (Department of Biochemistry, Okayama University of Science, Okayama, Japan) for the gift of soluble purified cytochrome b5 protein. Funding The present study was fully supported by the research grant of applied science to educate and direct graduate in both Chubu University and Fujita Health University. No additional financial support is not added to this present research focused on the enzyme function. Conflict of Interest None declared. References 1 D’Autréaux B., Toledano M.B. ( 2007) ROS as signaling molecules: mechanisms that generate specificity in ROS homeostasis. Nat. Rev. Mol. Cell Biol . 8, 813– 824 Google Scholar CrossRef Search ADS PubMed  2 Finkel T. ( 2011) Signal transduction by reactive oxygen species. J. Cell Biol.  194, 7– 15 Google Scholar CrossRef Search ADS PubMed  3 Turpaev K.T. ( 2002) Reactive oxygen species and regulation of gene expression. Biochemistry  67, 281– 292 Google Scholar PubMed  4 Bae Y.S., Oh H., Rhee S.G., Yoo Y.D. ( 2011) Regulation of reactive oxygen species generation in cell signaling. Mol. Cells  32, 491– 509 Google Scholar CrossRef Search ADS PubMed  5 Bánfi B., Molnár G., Maturana A., Steger K., Hegedûs B., Demaurex N., Krause K.-H. ( 2001) ACa2+-activated NADPH oxidase in testis, spleen, and lymph nodes. J. Biol. Chem . 276, 37594– 37601 Google Scholar CrossRef Search ADS PubMed  6 Bánfi B., Tirone F., Durussel I., Knisz J., Moskwa P., Molnár G.Z., Krause K.-H., Cox J.A. ( 2004) Mechanism of Ca2+ activation of the NADPH oxidase 5 (NOX5). J. Biol. Chem . 279, 18583– 18591 Google Scholar CrossRef Search ADS PubMed  7 Bánfi B., Malgrange B., Knisz J., Steger K., Dubois-Dauphin M., Krause K.-H. ( 2004) NOX3, a superoxide-generating NADPH oxidase of the inner ear. J. Biol. Chem . 279, 46065– 46072 Google Scholar CrossRef Search ADS PubMed  8 Nisimoto Y., Jackson M.H., Ogawa H., Kawahara T., Lambeth J.D. ( 2010) Constitutive NADPH dependent electron transferase activity of the Nox4 dehydrogenase domain. Biochemistry  49, 2433– 2442 Google Scholar CrossRef Search ADS PubMed  9 Nisimoto Y., Diebold A.B., Gomes C.D., Lambeth J.D. ( 2014) Nox4: a hydrogen peroxide-generating oxygen sensor. Biochemistry  53, 5111– 5120 Google Scholar CrossRef Search ADS PubMed  10 Ambasta R.K., Kumar P., Griendling K.K., Schmidt H.H., Busse R., Brandes R.P. ( 2004) Direct interaction of the novel Nox proteins with p22phox is required for the formation of a functionally active NADPH oxidase. J. Biol. Chem . 279, 45935– 45941 Google Scholar CrossRef Search ADS PubMed  11 Parkos C.A., Dinauer M.C., Walker L.E., Allen R.A., Jesaitis A.J., Orkin S.H. ( 1988) Primary structure and unique expression of the 22-kilodalton light chain of human neutrophil cytochrome b. Proc. Natl. Acad. Sci. USA  85, 3319– 3323 Google Scholar CrossRef Search ADS   12 Zhu Y., Marchal C.C., Cashbon A.J., Stull N., von Lohneysen K., Kraus U.C., Jesaitis A.J., McCmick S., Nauseef W.M., Dinauer M.C. ( 2006) Deletion of p22phox subunit of flavocytochrome b558: identification of regions critical for gp91phox maturation and NADPH oxidase activity. J. Biol.Chem . 281, 30336– 30346 Google Scholar CrossRef Search ADS PubMed  13 Yu L., Zhen L., Dinauer M.C. ( 1997) Biosynthesis of the phagocyte NADPH oxidase cytochrome b558: role of heme incorporation and heterodimer formation in maturation and stability of gp91phox and p22phox subunits. J. Biol. Chem . 272, 27286– 27294 14 Nakano Y., Banfi B., Jesaitis A.J., Dinauer M.C., Allen L.-A.H., Nauseef W.M. ( 2007) Critical roles for p22phox in the structural maturation and subcellular targeting of Nox3. Biochem. J . 403, 97– 108 Google Scholar CrossRef Search ADS PubMed  15 Martyn K.D., Frederick L.M., von Loehneysen K., Dinauer M.C., Knaus U.G. ( 2006) Functional analysis of Nox4 reveals unique characteristics compared to other NADPH oxidase. Cell Signal  18, 69– 82 Google Scholar CrossRef Search ADS PubMed  16 Serrander L., Cartierr L., Bendard K., Banfi B., Lardy B., Plastre O., Sienkiewicz A., Forro L., Schlegel W., Krause K.-H. ( 2007) NOX4 activity is determined by mRNA levels and reveals a unique pattern of ROS generation. Biochem. J . 406, 105– 114 Google Scholar CrossRef Search ADS PubMed  17 Krause K.-H. ( 2004) Tissue distribution and putative physiological function of NOX family NADPH oxidase. Jpn. J. Infect. Dis . 57, S28– S29 Google Scholar PubMed  18 Lambeth J.D., Kawahara T., Diebold B. ( 2007) Regulation of Nox and Duox enzymatic activity and expression. Free Radical Biol. Med . 43, 319– 331 Google Scholar CrossRef Search ADS   19 Ago T., Kuroda J., Pain J., Fu C., Li H., Sadoshima J. ( 2010) Upregulation of Nox4 by hypertrophic stimuli promotes apoptosis and mitochondrial dysfunction in cardiac myocytes. Circ. Res . 106, 1– 12 Google Scholar CrossRef Search ADS   20 Takac I., Schröder K., Zhang L., Lardy B., Anilkuman N., Lambeth J.D., Shah A.M., Morel F., Brandes R.P. ( 2011) The E-loop is involved in hydrogen peroxide formation by the NADPH oxidase Nox4. J. Biol. Chem . 286, 13304– 13313 Google Scholar CrossRef Search ADS PubMed  21 Van Buul J.D., Fernandez-Borja M., Anthony E.C., Hordijk P.L. ( 2005) Expression and localization of Nox2 and Nox4 in primary human endothelial cells. Antioxidant Redox Signal . 7, 308– 317 Google Scholar CrossRef Search ADS   22 Hilenski L.L., Clempus R.E., Quinn M.T., Lambeth J.D., Griendling K.K. ( 2004) Distinct subcellular localizations of Nox1 and Nox4 in vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol . 24, 677– 683 Google Scholar CrossRef Search ADS PubMed  23 Geiszt M., Kopp J.B., Varnai P., Leto T.L. ( 2000) Identification of renox, an NAD(P)H oxidase in kidney. Proc. Natl. Acad. Sci. USA  97, 8010– 8014 Google Scholar CrossRef Search ADS   24 Jackson H.M., Kawahara T., Nisimoto Y., Smith S.M.E., Lambeth J.D. ( 2010) Nox4 B-loop creates an interface between the transmembrane and dehydrogenase domains. J. Biol. Chem . 285, 10281– 10290 Google Scholar CrossRef Search ADS PubMed  25 Nisimoto Y., Ogawa H., Miyano K., Tamura M. ( 2004) Activation of the flavoprotein domain of gp91phox upon interaction with N-terminal p67phox (1- 210) and the Rac complex. Biochemistry  43, 9567– 9575 Google Scholar CrossRef Search ADS PubMed  26 Lutter R., van Schaik M.L.J., van Zwieten R., Wever R., Roos D., Hamers M.N. ( 1985) Purification and partial characterization of the b-type cytochrome from human polymorphonuclear Leukocytes. J. Biol. Chem . 260, 2237– 2244 Google Scholar PubMed  27 Vermilion J.L., Ballou D.P., Massey V., Coon M.J. ( 1981) Separate roles for FMN and FAD in catalysis by liver microsomal NADPH-cytochrome P-450 reductase. J. Biol. Chem.  256, 266– 277 Google Scholar PubMed  28 Zhao B., Summers F.A., Mason R.P. ( 2012) Photooxidation of Amplex Red to resorufin: implications of exposing the Amplex Red assay to light. Free Radic. Biol. Med . 53, 1080– 1087 Google Scholar CrossRef Search ADS PubMed  29 Rodrigues J.V., Gomes C.M. ( 2010) Enhanced superoxide and hydrogen peroxide detection in biological assays. Free Radical Biol. Med . 49, 61– 66 Google Scholar CrossRef Search ADS   30 Kimura S., Nishida H., Iyanagi T. ( 2001) Effects of flavin-binding motif amino acid mutations in the NADH-cytochrme b5 reductase catalytic domain on protein stability and catalysis. J. Biochem . 130, 481– 490 Google Scholar CrossRef Search ADS PubMed  31 Cheng G., Cao Z., Xu X., Meir E.G.V., Lambeth J.D. ( 2001) Homologs of gp91phox: cloning and tissue expression of Nox3, Nox4, and Nox5. Gene  269, 131– 140 Google Scholar CrossRef Search ADS PubMed  32 Shiose A., Kuroda J., Tsuruya K., Hirai M., Hirakata H., Naito S., Hattori M., Sakaki Y., Sumimoto H. ( 2001) A novel superoxide-producing NAD(P)H oxidase in kidney. J. Biol. Chem.  276, 1417– 1423 Google Scholar CrossRef Search ADS PubMed  33 Lambeth J.D. ( 2004) NOX enzymes and the biology of reactive oxygen. Nat. Rev. Immunol . 4, 181 Google Scholar CrossRef Search ADS PubMed  34 Lambeth J.D., Cheng G., Arnold R.S., Edens W.E. ( 2000) Novel homologs of gp91phox. Trends Biochem. Sci . 25, 459– 461 Google Scholar CrossRef Search ADS PubMed  35 von Löhneysen K., Noack D., Jesaitis A.J., Dinauer M.C., Knaus U.G. ( 2008) Mutational analysis reveals distinct features of the Nox4-p22phox complex. J. Biol. Chem.  283, 35273– 35282 Google Scholar CrossRef Search ADS PubMed  36 Denu J.M., Tanner K.G. ( 1998) Specific and reversible inactivation of tyrosine phosphatase by hydrogen peroxide: evidence for a sulfenic acid intermediate and implications for redox regulation. Biochemistry  37, 5633– 5642 Google Scholar CrossRef Search ADS PubMed  37 Barrett W.C., DeGnore J.P., König S., Fales H.M., Keng Y.-F., Zhang Y., Yim M.B., Chock P.B. ( 1999) Regulation of PTP1B via glutathionylation of the active site cysteine. Biochemistry  38, 6699– 6705 Google Scholar CrossRef Search ADS PubMed  38 Ziegler D.M. ( 1985) Role of reversible oxidation-reduction of enzyme thiols-disulfides in metabolic regulation. Annu. Rev. Biochem.  54, 305– 329 Google Scholar CrossRef Search ADS PubMed  39 Gilbert H.F. ( 1995) Thiol/disulfide exchange equilibria and disulfide bond stability. Methods Enzymol . 251, 8– 28 Google Scholar CrossRef Search ADS PubMed  40 Jones D.P., Go Y.M., Anderson C.L., Ziegler T.R., Kinkade J.M., Kirlin W.G. ( 2004) Cysteine/cystine couple is a newly recognized node in the circuitry for biologic redox signaling and Control. Faseb J . 18, 1246– 1248 Google Scholar CrossRef Search ADS PubMed  41 Netto L.E.S., Oliveira M.A., Monteiro G., Demasi A.P.D., Cussiol J.R.R., Discola K.F., Demasi M., Silva G.M., Alves S.V., Faria V.G., Horta B.B. ( 2007) Reactive cysteine in proteins: protein folding, antioxidant defense, redox signaling and more. Comparative Biochem. Physiol. Part C  146, 180– 193 42 Go Y.M., Jones D.P. ( 2010) Redox control systems in the nucleus: mechanisms and functions. Antioxid. Redox Signal . 13, 489– 509 Google Scholar CrossRef Search ADS PubMed  43 Miki H., Funato Y. ( 2012) Regulation of intracellular signaling through cysteine oxidation by reactive oxygen species. J. Biochem . 151, 255– 261 Google Scholar CrossRef Search ADS PubMed  44 Haddad J.J. ( 2002) Science review: redox and oxygen-sensitive transcription factors in the regulation of oxidant-mediated lung injury: role for nuclear factor-kB. Crit. Care  6, 481– 490 Google Scholar CrossRef Search ADS PubMed  45 Merchant A.A., Singh A., Matsui W., Biswal S. ( 2011) The redox-sensitive transcription factor Nrf2 regulates murine hematopoietic stem cell survival independently of ROS levels. Blood  118, 6572– 6579 Google Scholar CrossRef Search ADS PubMed  46 Rhee S.G. ( 1999) Redox signaling: hydrogen peroxide as intracellular messenger. Exp. Mol. Med . 31, 53– 59 Google Scholar CrossRef Search ADS PubMed  47 Stone J.R., Yang S. ( 2006) Hydrogen peroxide: a signaling messenger. Antioxid. Redox Signal  8, 243– 270 Google Scholar CrossRef Search ADS PubMed  48 Jackson H.M. ( 2010) Structure and Functional Analysis of NADPH Oxidase 4. Ph. D. thesis. Emory University, ATL, pp. 1– 210 49 Segal A.W., West I., Wientjes F.B., Nugent J.H.A., Chavan A.J., Haley B., Garcia R.C., Rosen H., Scrace G. ( 1992) Cytochrome b-245 is a flavocytochrome containing FAD and the NADPH-binding site of the microbicidal oxidase of phagocytes. Biochem. J.  284, 781– 788 Google Scholar CrossRef Search ADS PubMed  50 Taylor W.R., Jones D.T., Segal A.W. ( 1993) A structural model for the nucleotide binding domains of the flavocytochrome b-245 beta-chain. Protein Sci . 2, 1675– 1685 Google Scholar CrossRef Search ADS PubMed  Abbreviations Abbreviations DH dehydrogenase DPI diphenylene iodonium Duox dual oxidase GRx glutaredoxin HEK human embryonic kidney HRP horseradish peroxidase MBP maltose binding protein NEF nucleus-enriched fraction Nox NADPH oxidase PMSF phenylmethanesulfonyl fluoride phox phagocite oxidase ROS reactive oxygen species SOD superoxide dismutase TM transmembrane TRx thioredoxin © The Author(s) 2018. Published by Oxford University Press on behalf of the Japanese Biochemical Society. All rights reserved This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices) http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png The Journal of Biochemistry Oxford University Press

NADPH oxidase 4 function as a hydrogen peroxide sensor

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Oxford University Press
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© The Author(s) 2018. Published by Oxford University Press on behalf of the Japanese Biochemical Society. All rights reserved
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0021-924X
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1756-2651
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10.1093/jb/mvy014
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Abstract

Abstract Nox4, a member of the NADPH- and oxygen-dependent oxidoreductases that generate reactive oxygen species (ROS), is widely expressed and constitutively active. To understand better its function and regulation, specific mutations in the Nox4 dehydrogenase (DH) domain were examined for effects on Nox4 oxidase activity. Transfection of His6-tagged Nox4 increased the amount of p22phox subunit in HEK293 cells, and a higher level of the heterodimer was observed in the nucleus-enriched fraction (NEF). NEF from Nox4-expressing HEK293 cells exhibited oxygen and H2O2 concentration-dependent NADPH oxidation rate. In Nox4-expressing cells, NEF and its partially purified form, the Nox4(P437H) mutant almost completely lost its oxidase activity, while Nox4(C546S), Nox4(C546L) or/and (C547L) had a significantly decreased rate of ROS production. The NADPH-dependent reduction of cytochrome c or cytochrome b5 by purified Nox4 DH domain was found regulated by the H2O2 concentration, and C546L and C547L mutants showed lower rates of the hemeprotein reduction. These conserved Cys residues in the DH domain respond to the cytosolic H2O2 concentration to regulate Nox4 activity. electron transfer, flavocytochrome, NADPH oxidase, reactive oxygen, redox regulation Nox enzymes comprise a family of NADPH oxidoreductases that generate reactive oxygen species (ROS) in a variety of cell types and tissues in response to numerous (both normal and abnormal) physiological signals (1, 2). The seven Nox isoforms (Nox1-5, Duox1 and Duox2) expressed in human tissues can be distinguished by their tissue distribution, structure and regulation, and reactive oxygen product. However, all seven Nox isoform proteins consist of highly conserved structural features as flavocytochrome; the C-terminal dehydrogenase (DH) domain contains binding sites for FAD and NADPH. The N-terminal transmembrane (TM) region consists of a six α-helical structure that involves four conserved histidine residues, located in the third and fifth TM helices that co-ordinate two heme groups (heme bL and bH). Regulated ROS production occurs at considerably reduced levels in most non-phagocytic tissues compared to activated phagocytes, or neutrophils (3, 4). Non-phagocytic Nox1 and Nox3 and phagocytic Nox2 form heterodimers with p22phox in membranes and require additional cytosolic regulatory proteins for function. Nox4 associates with p22phox in membranes but does not require cytosolic proteins. Nox5 and the dual oxidases Duox1 and Duox2 are activated by calcium (Ca2+) through a cytosolic EF-hand containing Ca2+ binding domain (5, 6). Nox1-3 and Nox5 generate primarily superoxide; Nox4, Duox1 and Duox2 produce mainly H2O2 (7–9). Nox4 may be unique among the Nox isoforms. Though Nox4 associates with p22phox in membranes, it does not require additional regulatory proteins or the proline-rich domain in p22phox, which is responsible for binding to the regulatory proteins, for activity (10, 11). Mutations in p22phox that inhibit Nox1-3 catalytic function do not alter Nox4-p22phox oxidase activity (12–14), suggesting that p22phox is required for Nox4 expression and specific membrane localization (15) but not for catalytic function. This has led to the concept that Nox4-dependent ROS generation is regulated primarily by its expression level (16–18). Moreover, most Nox/Duox isoforms are detected in many kinds of cells to produce ROS, and Nox4 is mainly observed in the membranes of endoplasmic reticulum, nucleus and mitochondria (19–22). In addition, unlike the other six isoforms, Nox4 is constitutively active (15, 23). Electron transfer activities of Nox1-5 DH domain proteins, Nox2 TM-Nox4 DH and Nox4 TM-Nox2 DH chimaera proteins were compared and only the Nox4 DH protein showed a significantly high rate without added activating subunits (8, 24), while Nox2 DH was activated by the association with p67phox-Rac1 complex (25). The DH domain of only Nox4 is constitutively ‘turned on’ compared with the DH domains of Nox1-3 and Nox5, thus, this property of the DH domain accounts for the constitutive active of the Nox4 holoenzyme (8). Here, to understand the mechanism of Nox4 regulation, mutations in the DH domain were examined for effects on oxidase activity. The present study suggested that H2O2-sensitive Nox4 DH domain containing cysteine residue(s) plays an important role in regulating ROS production in non-phagocytic cells. Experimental Procedures Materials Full-length cDNA encoding human Nox4 (amino acid residues 1-578) and N-terminally His6-tagged Nox4 cDNA were cloned into pcDNA3.1 and pCIG vectors, respectively (Invitrogen). cDNA encoding Nox-DH domains (Nox2 and Nox4) were subcloned into pMAL-C2X (New England Biolabs). DH domains corresponded to residues 283-570 for Nox2 and 304-578 for Nox4. These constructs, producing maltose binding protein (MBP)-tagged C-terminal Nox2 and Nox4 homologs, were expressed in Escherichia coli (E. coli) BL-21. The goat anti-rabbit IgG secondary anti-body linked to horseradish peroxidase (HRP) and pre-stained molecular weight markers for SDS-PAGE were from Bio-Rad. The polyclonal anti-body to a C-terminal peptide (residues 500–578) of Nox4 was from Novus Biologicals Inc., and monoclonal anti-bodies to MBP, amylose agarose and Factor Xa protease were from New England Biolabs. Mouse monoclonal anti-body 44.1 against human p22phox was from Santa Cruz Biotech, and mouse monoclonal anti-body #2366 to His6 was from Cell Signalling Technologies. The goat anti-mouse secondary anti-bodies linked to HRP came from Promega. Protease inhibitor cocktail (EDTA-free) and Amplex Red were from Roche and Invitrogen, respectively. Ferricytochrome c, glucose 6-phosphate DH (G6PDH), FAD, NADPH, NADP+, diphenylene iodonium (DPI), phenylmethanesulfonyl fluoride (PMSF), protein A-agarose Fast flow [50% (v/v)], 3, 3’-diaminobenzidine and nuclei EZ lysis buffer were purchased from Sigma Aldrich (St. Louis, MO). Site-directed mutagenesis and transient transfection of Nox4 According to the previously described methods (13), His6-tagged Nox4 point mutations Pro-437 mutated to His (denoted P437H), Cys-546 to Ser (C546S), Cys-546 to Leu (C546L) and Cys-547 to Leu (C547L) were generated using site-directed mutagenesis with human full length Nox4 cDNA. HEK293 cells were seeded at 1 x 106 cells/plate (10 cm diameter) and grown 24 h to 40–50% confluence in Dulbecco’s modified Eagle’s medium with 10% foetal bovine serum, 100 units/ml penicillin and 0.1 mg/ml streptomycin. Cells were transfected 48 h prior to use with mammalian expression vectors encoding His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4 (C546L/C547L) or empty vector, using FuGENE6 (Roche Molecular Biochemicals). Separation of nucleus-enriched fraction Transiently transfected HEK293 cells (3.5 ∼ 5 x 108 cells) cultured in several 10 cm tissue culture plates were harvested, washed twice with PBS, suspended in nuclei EZ lysis buffer, pH 7.4, containing 25 μM FAD and protease inhibitor cocktail (Complete Mini, Roche Diagnostics) plus 0.2 mM PMSF and disrupted using a glass homogenizer with a loose fitting pestle (3 min at 4°C). The homogenate was centrifuged at 800 x g, for 5 min in a KUBOTA 5922 rotor at 4°C to collect a nucleus-enriched fraction (NEF). The pooled NEF (30 μg) was subjected to 5–20% Tris-Glycine SDS-PAGE, electrotransferred to PVDF membranes (Millipore) and confirmed to be nucleus-enriched by immunoblotting with an anti-body to Lamin A. This fraction was also evaluated for oxygen or H2O2 concentration-dependent NADPH oxidase activity using fluorescence or absorption change. Western blot analysis His6-tagged Nox4 protein located in NEF of HEK293 cells was detected by Western blotting. Nucleus-enriched pellet extracts were prepared using the Nonidet P-40 lysis buffer containing 10% glycerol as described previously (9). Extracted proteins were separated on SDS-PAGE (5–20% gel) and transferred to an Immobilon PVDF membrane (Millipore). Proteins were visualized by incubation with primary anti-bodies overnight at 4°C with gentle shaking, and then with horseradish-linked secondary anti-body (1:3000 dilution, 2 h). Bands were detected by Luminescent Image Analyzer (Image Quant LAS 4000) after addition of Super Signal West Pico Chemiluminescent Substrate (Thermo Scientific), according to the manufacturer’s instructions. Purified MBP-tagged Nox2 DH, Nox4 DH, Nox4 DH (P437H), Nox4 DH (C546L) and Nox4 DH (C547L) proteins were subjected to 5–20% SDS-PAGE followed by immunoblotting using an anti-MBP anti-body and visualized as described above. Spectrophotometric measurement of Heme in NEF The NEF extracts were prepared using the Nonidet P-40 buffer according to the previously described method (9). The extracts prepared from HEK293 cells transfected with vector alone, vector encoding His6-Nox4, His6-Nox4(C546L), His6-Nox4 (C547L) or His6-Nox4(C546L/C547L) were used for the quantification of heme. Reduced minus oxidized difference spectra were recorded at 10 min intervals after addition of a few crystals of sodium dithionite until a stable spectrum was obtained. A molar absorption coefficient (ε426–408 nm) of 200 mM−1.cm−1 (26) for the Soret band was used for calculations. Superoxide generating activity Superoxide generation was assessed using superoxide dismutase (SOD)-inhibitable ferricytochrome c reduction quantified at 550 nm using an extinction coefficient of 21.1 mM−1.cm−1 (27). The indicated amount of HEK293 cell number, NEF or isolated Nox4 DH domain protein was added to the assay buffer [25 mM Hepes (pH 7.3) containing 25 μM FAD, 80 μM ferricytochrome c, 0.12 M NaCl, 3 mM KCl and 1 mM MgCl2], after which 50 μM NADPH was added and cytochrome c reduction was monitored in the presence and absence of 200 units of SOD at 25°C. Measurement of hydrogen peroxide generation Amplex Red (100 μM) plus 1.5 units/ml HRP was included in the assay mixture, and the reaction was initiated by the addition of intact cells, NEF or partially purified His6-Nox4 protein. The linear increase in the fluorescence of resorufin produced by oxidation of Amplex Red was measured (28). For experiments monitoring fluorescence, excitation and emission wavelengths of 572 nm and 583 nm, respectively, were used for microplate measurements. Reactions were monitored at 25°C for 10 min using Mithras LB940 multimode microplate reader (BERTHOLD Technologies). For NEF, 0.1 mg/ml final protein concentration was added to a reaction mixture containing 25 μM FAD, 50 μM glucose 6-phosphate, 50 μM NADP+ and commercial G6PDH (0.25 U/ml) was included. We used an NADPH-generating assay system consisting of glucose 6-phosphate, G6PDH and NADP+ to markedly decrease the background rate of Amplex Red oxidation by reduced pyridine nucleotide. The remaining low residual rate of enzyme-independent Amplex Red oxidation was then subtracted to obtain correct rates. The concentration of Amplex Red oxidized was calculated using an extinction coefficient of 54 mM−1•cm−1 at 572 nm (29), or when fluorescence was measured, using a standard curve generated from the addition of known amounts of hydrogen peroxide. Expression of Nox2 DH and Nox4 DH domains Truncated Nox DH clones were obtained by PCR using Nox cDNA cloned in the pMAL-C2X plasmid as the template. According to previously described methods (8, 24), PCR products for truncated Nox2 DH (residues 283-570), Nox4-DH (residues 304-578) and Nox4 DH mutant (residues 304-578, P437H, C546L or C547L) domains were purified using a PCR purification kit (Qiagen). The purified DNA fragments were ligated into the BamHI and HindIII restriction sites for the Nox2 DH domain, and BamHI and SalI for Nox4 DH domain in pMAL-C2X vector and transformed into E. coli. Transformants were selected from LB/ampicillin plates, and plasmids were isolated from 2 ml cultures as described previously (8). The plasmids were digested with restriction enzymes and separated on 1% agarose to confirm the presence of the insert. DNA sequences of the four clones were confirmed by nucleotide sequencing. Purification of expressed His6-Nox4 and MBP-Nox DH proteins His6-Nox4 or His6-Nox4 mutant was expressed in HEK293 cells and its partial purification was performed by a Ni-NTA affinity column chromatography as described previously (9). MBP-DH fusion proteins were induced in E. coli at 37°C by addition of 0.1 mM IPTG for 2.5 h and frozen at –80°C. Thawed cells were sonicated (3 x 10 s) and solubilized in 50 mM Hepes buffer, pH 7.5, containing 0.5 M NaCl, 1 mM PMSF, 1 mM EDTA, 1 mM dithiothreitol, protease inhibitor cocktail (1 μg/ml) and 0.15 M L-arginine at 3°C. Purification was performed by amylose-agarose column chromatography (10 x 15 mm) according to the methods described previously (8, 9). MBP-depletion was performed in the presence of Factor Xa for 6 h at 4°C according to the manufacturer’s instructions (New England Biolabs). MBP-fused and depleted proteins were used in 1 week to avoid gradual proteolysis and loss of activity. Purified MBP-fused and deleted Nox2-DH and Nox4 DH proteins were subjected to 5–20% SDS-PAGE followed by protein staining using Coomassie brilliant blue. Pyridine nucleotide-dependent electron transfer activity NADPH-dependent cytochrome c and cytochrome b5 reductase activities were assayed at 25°C according to the previously described methods (8). The activities were assayed in 1 ml volume of assay buffer [25 mM Hepes, pH 7.3, containing 0.12 M NaCl, 3 mM KCl, 1 mM MgCl2, 25 μM FAD, protease inhibitor cocktail (1 μg) and 80 μM electron acceptor]. After the mixture that included the purified Nox4 DH protein in the presence and absence of H2O2 had been pre-incubated for 5 min, the reaction was initiated by the addition of 50 μM NADPH. The reduction rates of electron acceptors were quantified by monitoring the absorbance changes at the appropriate wavelengths, and millimolar extinction coefficients for cytochrome c (21.1 mM−1•cm−1) (27) and cytochrome b5 (19 mM−1•cm−1) (30) at 556 nm were used to calculate the quantity of each electron acceptor reduced. Spectrophotometric measurements were performed using a Hitachi spectrophotometer with a temperature-controlled cuvette compartment. Oxygen concentration-dependent H2O2 generation by NEF The gas equilibration system consisted of a tightly capped 1.5 ml cuvette with a gas delivery needle and a gas exit needle that also served as a delivery port for addition of reagents. ROS measurement was conducted at 20°C in a total volume of 0.8 ml as described (9). The reaction mixture was equilibrated by gently bubbling (approximately one bubble per second) with the indicated percent of oxygen/nitrogen gas mixtures, with continuous gentle stirring for 10 min using a magnetic stirrer. Fluorescence increase at 583 nm due to Amplex Red oxidation was measured as described above. NADPH oxidation NADPH oxidation was measured spectrophotometrically at 340 nm in the presence of either NEF separated from HEK293 cells or purified MBP-Nox4 DH protein in 1 ml of the assay buffer as described above. After the assay mixture containing the sample was pre-incubated at 20°C for 5 min in the presence or absence of H2O2, the reaction was started by adding 50 μM NADPH. The amount of NADPH oxidized for 10 min was calculated using a molar absorption coefficient of 6.24 mM−1.cm−1 at 340 nm. The NADPH oxidation rate was also measured by the fluorescence decrease at 460 nm when excited at 340 nm. Results Analysis of C-terminal DH domain mutation of His6-Nox4 expressed in HEK293 cell To determine the importance of the NADPH DH domain in constitutive active Nox4 enzyme, point substitutions of proline residue at 437 and cysteine residue at 546 or/and 547 were generated by site-directed mutagenesis. Mutant His6-tagged Nox4 was expressed in HEK293 cells, and ROS-generating activity was monitored. Mutation of P437H resulted in nearly complete loss of Nox4 activity, whereas the C546S, C546L or/and C547L mutation indicated a remarkably decreased activity of wild-type Nox4 (Fig. 1A). Reactive oxygen product generation of His6-tagged Nox4 and mutant Nox4 (C546S, C546L or/and C547L) was verified to be largely inhibited by the general flavoprotein DH inhibitor, DPI. ROS generation was mainly detected in wild-type Nox4- and Nox4 mutant (C546S, C546L or/and C547L)-expressing cells, with the H2O2 identity validated due to inhibition by catalase and superoxide generating activity was greatly lower (Fig. 1B and C). The substitution of cysteine-546 to serine or leucine resulted in a significantly lower H2O2 and O2- producing activities as compared with wild-type Nox4 (Fig. 1B and C), and the mutation of either C547L or both C546L and C547L also showed a further decrease of ROS generating activity as observed in Fig. 1A–C, respectively. Consistent with His6-tagged Nox4, intact HEK293 cells stably expressing Nox4 also indicated that the major product was H2O2 and 40% or less superoxide was observed (9). These data also support that whole cells from His6-Nox4 transfected HEK293 cells were observed to generate mainly H2O2 at nearly the same rate as intact cells stably expressing native Nox4. Fig. 1 View largeDownload slide ROS generation by Nox4 in HEK293 cells—cDNA encoding His6-Nox4 (wild type), His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or empty vector alone (Mock) was transfected in HEK293 intact cells, as described in Experimental Procedures. (A) ROS was measured in the absence (white bars) and presence (grey bars) of 20 μM DPI using luminol chemiluminescence with 20 μM luminol and 0.32 unit HRP in 200 μl total volume. Results are shown as the mean ± SD. of three separate wells, and are representative of three separate transfection tests. (B) Hydrogen peroxide production was measured in intact His6-Nox4-expressing cells. The reaction was initiated by adding 5.2 x 105 cells to 0.8 ml of 25 mM Hepes, pH 7.3, containing 1.5 unit HRP and 100 μM Amplex Red. Amplex Red oxidation due to H2O2 was measured as the fluorescence increase at 583 nm when excited at 572 nm in the absence (white bars) and presence of 50 unit/ml of catalase (black bars). (C) Superoxide generation was quantified as superoxide dismutase (SOD)-inhibitable cytochrome c reduction at 550 nm, carried out in 1 ml of assay buffer, pH 7.3, containing 80 μM ferricytochrome c with or without SOD (final 300 U/ml) as described in Experimental Procedures. Values represent the mean activity ± SEM of three determinations using one set of transfected cells, and are representative of three transfection experiments. Fig. 1 View largeDownload slide ROS generation by Nox4 in HEK293 cells—cDNA encoding His6-Nox4 (wild type), His6-Nox4(P437H), His6-Nox4(C546S), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or empty vector alone (Mock) was transfected in HEK293 intact cells, as described in Experimental Procedures. (A) ROS was measured in the absence (white bars) and presence (grey bars) of 20 μM DPI using luminol chemiluminescence with 20 μM luminol and 0.32 unit HRP in 200 μl total volume. Results are shown as the mean ± SD. of three separate wells, and are representative of three separate transfection tests. (B) Hydrogen peroxide production was measured in intact His6-Nox4-expressing cells. The reaction was initiated by adding 5.2 x 105 cells to 0.8 ml of 25 mM Hepes, pH 7.3, containing 1.5 unit HRP and 100 μM Amplex Red. Amplex Red oxidation due to H2O2 was measured as the fluorescence increase at 583 nm when excited at 572 nm in the absence (white bars) and presence of 50 unit/ml of catalase (black bars). (C) Superoxide generation was quantified as superoxide dismutase (SOD)-inhibitable cytochrome c reduction at 550 nm, carried out in 1 ml of assay buffer, pH 7.3, containing 80 μM ferricytochrome c with or without SOD (final 300 U/ml) as described in Experimental Procedures. Values represent the mean activity ± SEM of three determinations using one set of transfected cells, and are representative of three transfection experiments. The expression of Nox4 and its mutant proteins were examined by expressing His6-tagged Nox4 in HEK293 cells, followed by Western blotting (Fig. 2A). To determine if Nox4 and p22phox formed active heterodimers in NEF, we assessed the impact of p22phox expression on constitutively active Nox4. In NEF low levels of endogenous Nox4 and p22phox subunits are observed, but further expression of Nox4 subunit enhanced the steady-state level of p22phox. The results suggest that the transfection of Nox4 increased the amount of p22phox protein in HEK293 cells, and a higher level of the heterodimer was found in NEF. Almost similar levels of expression were detected for the four mutants compared with wild-type Nox4. Nox4 heterodimerizes with p22phox, and this association is assumed to be required for stability and location in the membrane of the nucleus. In accord with previous report (9), the increased Nox4 protein and ROS production were clearly confirmed in the NEF prepared from Nox4-expressing cells. Therefore, Nox4-enriched NEF was used in the present studies. Fig. 2 View largeDownload slide Determination of ROS level produced in a NEF or partially purified His6-Nox4 from HEK293 cells—(A) Expression of Nox4 and p22phox were determined in immunoblots of nuclear enriched fractions (NEF) (30 µg) prepared from HEK293 cells transfected with His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or Mock-transfected. (B) The NEF (0.1 mg) was supplemented with FAD, NADP+, glucose 6-phosphate and glucose 6-phosphate DH in NADPH-generating system, and hydrogen peroxide producing activity (white bars) was monitored by Amplex Red oxidation as described in Fig. 1B. Superoxide generation was assayed by SOD-inhibitable cytochrome c reduction monitored at 550 nm (grey bars). (C) NADPH-dependent H2O2 production of NEF was monitored in the absence (white bars) and presence (grey bars) of 20 μM DPI by measuring Amplex Red fluorescence change at 583 nm when excited at 572 nm. (D) Reduced minus oxidized difference spectra of NEF prepared from HEK293 cells. NEF expressed Mock (2.51 mg/ml), His6-Nox4(Wild) (2.05 mg/ml), His6-Nox4(C546L) (2.28 mg/ml) or His6-Nox4(C547L) (2.30 mg/ml) was reduced with a trace amount of Na2S2O4 for 5 min and difference spectra were measured at 20°C. (E) Partially purified His6-tagged forms (2.5 μg each) of Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L) were loaded onto SDS-PAGE gel (5–20%) and stained with Coomassie brilliant blue (top). NADPH-dependent H2O2 generating activity of purified His6-Nox4 was measured in the absence (white bars) and presence of either 20 μM DPI (grey bars) or 50 unit/ml of catalase (black bars) by monitoring Amplex Red fluorescence change at 583 nm as described above. Values represent the mean activity ± SEM of three determinations using one set of NEF (B, C) and purified protein (E). Similar results were obtained from three separate transfections for all experiments. Fig. 2 View largeDownload slide Determination of ROS level produced in a NEF or partially purified His6-Nox4 from HEK293 cells—(A) Expression of Nox4 and p22phox were determined in immunoblots of nuclear enriched fractions (NEF) (30 µg) prepared from HEK293 cells transfected with His6-Nox4, His6-Nox4(P437H), His6-Nox4(C546L), His6-Nox4(C547L), His6-Nox4(C546L/C547L) or Mock-transfected. (B) The NEF (0.1 mg) was supplemented with FAD, NADP+, glucose 6-phosphate and glucose 6-phosphate DH in NADPH-generating system, and hydrogen peroxide producing activity (white bars) was monitored by Amplex Red oxidation as described in Fig. 1B. Superoxide generation was assayed by SOD-inhibitable cytochrome c reduction monitored at 550 nm (grey bars). (C) NADPH-dependent H2O2 production of NEF was monitored in the absence (white bars) and presence (grey bars) of 20 μM DPI by measuring Amplex Red fluorescence change at 583 nm when excited at 572 nm. (D) Reduced minus oxidized difference spectra of NEF prepared from HEK293 cells. NEF expressed Mock (2.51 mg/ml), His6-Nox4(Wild) (2.05 mg/ml), His6-Nox4(C546L) (2.28 mg/ml) or His6-Nox4(C547L) (2.30 mg/ml) was reduced with a trace amount of Na2S2O4 for 5 min and difference spectra were measured at 20°C. (E) Partially purified His6-tagged forms (2.5 μg each) of Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L) were loaded onto SDS-PAGE gel (5–20%) and stained with Coomassie brilliant blue (top). NADPH-dependent H2O2 generating activity of purified His6-Nox4 was measured in the absence (white bars) and presence of either 20 μM DPI (grey bars) or 50 unit/ml of catalase (black bars) by monitoring Amplex Red fluorescence change at 583 nm as described above. Values represent the mean activity ± SEM of three determinations using one set of NEF (B, C) and purified protein (E). Similar results were obtained from three separate transfections for all experiments. In Fig. 2B, superoxide and hydrogen peroxide generation were compared in NEF from wild-type Nox4 and mutant Nox4-expressed cells. In the nuclear pellet fraction expressing Nox4, approximately 70% of ROS product was detected as H2O2, whereas 30% or less was observed as superoxide. Consistent with the whole cell assay as described in Fig. 1A and B, NEF expressing Nox4(P437H) almost completely lost H2O2 generation, and Nox4(C546L), Nox4(C547L) and Nox4(C546L/C547L) mutant released approximately 75%, 60% and 50% of H2O2, respectively, detected by wild-type Nox4 as the major ROS product. Inclusion of 20 μM DPI inhibited H2O2 generation more than 70% of wild-type Nox4 and Nox4(C546L, C547L or C546L/C547L) mutants (Fig. 2C). The concentration of heme in NEF prepared from non-transfected, His6-tagged Nox4, His6-Nox4(C546L), or His6-Nox4(C547L)-transfected HEK293 cells was 3.98, 10.9, 9.55 and 9.51 pmol/mg, respectively, indicating that the heme of wild-type His6-Nox4 and mutant Nox4(C546L or C547L) expressed in NEF was nearly the same level (Fig. 2D). In addition, the heme concentration in NEF from His6-Nox4(P437H)- or His6-Nox4(C546L/C547L)-transfected HEK293 was 10.5 and 9.63 pmol/mg protein, respectively. Therefore, the decrease in activity displayed by the Nox4 mutants was not due to altered protein expression, processing, or complex formation with p22phox. As shown in Fig. 2E, hydrogen peroxide generation of the partially purified His6-Nox4 was also compared in Nox4, Nox4(P437H), Nox4(C546L) and Nox4(C547L). In accord with the results obtained by using NEF, purified Nox4(P437H) mutant almost completely lost H2O2 generating activity and the individual C546L and C547L mutations considerably decreased the oxidase activity. In addition, Nox4(C546L/C547L) double mutant showed a little higher inhibitory effect than either C546L or C547L mutant form (data not shown). These results suggest a possible critical role of proline 437, cysteine 546 and cysteine 547 residues in the Nox4 DH domain for H2O2 generation. Effect of oxygen or hydrogen peroxide on NADPH oxidase turnover in Nox4-expressed nucleus- enriched fraction of HEK293 cells In the previous study (9), we reported that Nox4 activity is regulated not only by its expression level, but also by oxygen availability, and that it therefore functions as an oxygen sensor. Nuclear membrane-associated Nox4 has a high Km value for oxygen that allows it to respond to physiological wide ranges of oxygen concentration, such that its enzymatic activity must be linked to an effect on signalling that can be translated into a cellular response. In fact, Nox2-dependent superoxide generation in intact human neutrophils shows a Km for oxygen of about 3%; on the other hand, Nox4 in intact cells indicated a higher oxygen Km value for H2O2 generation of around 15%, corresponding to the Km range seen for other known oxygen sensing enzymes (9). Here we found that Nox4(C546L or C547L) mutants have a higher Km value for oxygen of about 23%, as compared with a Km of 17% detected in Nox4-expressed NEF (Fig. 3A). Additionally, NADPH oxidation in Nox4-expressing NEF showed a higher rate compared to control NEF (Fig. 3B). The presence of 50 μM H2O2 significantly decreased the rate of NADPH oxidation in Nox4-expressed NEF, and the addition of 200 unit SOD also slightly reduced the oxidation rate. Consistent with these data, a decrease in the hydrogen peroxide concentration-dependent NADPH oxidation rate was detected in NEF prepared from Nox4 and Nox4-mutant (C546L or C547L) expressing HEK293 cells (Fig. 3C and D). NADPH oxidation was determined by measuring absorption and fluorescence changes of NADPH, which are thought to correspond to the H2O2 concentration-dependent effect on the rate of NADPH oxidation. In fact, the NADPH oxidation of NEF from wild-type Nox4-transfected cells was a little more remarkably decreased than mutant forms (C546L or C547L) in the presence of a lower range of H2O2 concentrations (0–100 μM). In the presence of 50 μM H2O2, NADPH oxidase activity of NEF expressed wild-type Nox4 was approximately 52% decrease, while Nox4 (C546L) or (C547L) mutant indicated almost same decrease, 45 or 44%, respectively. Thus, the catalytic function of NADPH-dependent Nox4 enzyme appears to be closely regulated within the physiological range of cytosolic H2O2 concentrations as well as cellular pO2 values. Fig. 3 View largeDownload slide Hydrogen peroxide and oxygen concentration-dependent NADPH oxidation activity of NEF expressing His6-Nox4—(A) Oxygen concentration-dependent H2O2 production by NEF stably expressing His6-Nox4 (white circles), His6-Nox4(C546L) (filled circles) or His6-Nox4(C547L) (white triangles) was measured using Amplex Red fluorescence at 583 nm. A double-reciprocal plot of initial velocity versus O2 concentration on the top was formed to calculate Km and Vmax values. (B) After the NEF was preincubated for 5 min at 20°C in the presence of either H2O2 or SOD, 50 μM NADPH was added. The NADPH oxidation rate was monitored by measuring its absorption decrease at 340 nm without (white bars) or with added either 50 μM H2O2 (grey bars) or 200 U/ml SOD (filled bars). The values shown in (A) and (B) are the mean ± SEM of three determinations from one set of transfections, and is representative of three experiments repeated using three different sets of transfected cells. (C) NEF (78.5 μg) prepared from His6-Nox4- (white circles), His6-Nox4(C546L)- (filled circles) or His6-Nox4(C547L)-expressed cells (white triangles) was suspended in 0.8 ml of 25 mM Hepes, pH 7.3, containing 1 mM MgCl2 and the indicated concentrations of H2O2. After the NEF suspension was preincubated for 5 min at 20°C, 50 μM NADPH was added and measured the absorption change at 340 nm. (D) The rate of NADPH oxidation was monitored by fluorescence change at 460 nm when excited at 340 nm. The each symbol indicates the same NEF prepared from HEK293 as described in Fig. 3C. Data points and error bars in (C) and (D) indicate the mean ± SEM of three determinations from single assays, and the experiments shown are representative of three different experiments. Fig. 3 View largeDownload slide Hydrogen peroxide and oxygen concentration-dependent NADPH oxidation activity of NEF expressing His6-Nox4—(A) Oxygen concentration-dependent H2O2 production by NEF stably expressing His6-Nox4 (white circles), His6-Nox4(C546L) (filled circles) or His6-Nox4(C547L) (white triangles) was measured using Amplex Red fluorescence at 583 nm. A double-reciprocal plot of initial velocity versus O2 concentration on the top was formed to calculate Km and Vmax values. (B) After the NEF was preincubated for 5 min at 20°C in the presence of either H2O2 or SOD, 50 μM NADPH was added. The NADPH oxidation rate was monitored by measuring its absorption decrease at 340 nm without (white bars) or with added either 50 μM H2O2 (grey bars) or 200 U/ml SOD (filled bars). The values shown in (A) and (B) are the mean ± SEM of three determinations from one set of transfections, and is representative of three experiments repeated using three different sets of transfected cells. (C) NEF (78.5 μg) prepared from His6-Nox4- (white circles), His6-Nox4(C546L)- (filled circles) or His6-Nox4(C547L)-expressed cells (white triangles) was suspended in 0.8 ml of 25 mM Hepes, pH 7.3, containing 1 mM MgCl2 and the indicated concentrations of H2O2. After the NEF suspension was preincubated for 5 min at 20°C, 50 μM NADPH was added and measured the absorption change at 340 nm. (D) The rate of NADPH oxidation was monitored by fluorescence change at 460 nm when excited at 340 nm. The each symbol indicates the same NEF prepared from HEK293 as described in Fig. 3C. Data points and error bars in (C) and (D) indicate the mean ± SEM of three determinations from single assays, and the experiments shown are representative of three different experiments. Hydrogen peroxide concentration-dependent electron transferase activity of constitutive active Nox4 DH domain Herein, we explore the hypothesis that the structural features necessary for constitutively active Nox4 reside in a cytosol-facing DH domain that contains a binding site for NADPH and one for FAD. To determine whether this domain was sufficient to exhibit spontaneous electron transferase, the Nox4 DH domain was expressed and purified as an MBP fusion protein and electron transfer activity toward cytochrome c or cytochrome b5 was measured (Fig. 4). In addition, to investigate the structural features necessary for constitutive activity are ascribed to the Nox4 DH domain, Nox2 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domains were also expressed and purified to compare their NADPH-dependent electron transfer activities. Purified MBP-Nox DH domain fusion and MBP-depleted Nox DH proteins corresponded in size to their predicted molecular masses on SDS-PAGE (Fig. 4A-a, -b) and MBP-fused Nox DH proteins were recognized on Western blots using an anti-MBP anti-body (Fig. 4A–c). As previously reported (8), the MBP-Nox4 DH domain fusion protein showed significant NADPH-dependent electron transfer activity toward one-electron acceptor heme proteins (Fig. 4B). The MBP-Nox4 DH(C546L or C547L) mutant proteins showed a distinctive decrease in NADPH-dependent electron transferase activity, and the turnover number toward each electron acceptor was about 50–60% level observed for the wild-type MBP-Nox4 DH. In addition, Nox2 DH exhibited a negligible NADPH-dependent electron transferase activity, and the electron transfer rate of Nox4 DH(P437H) domain was very low for each electron acceptor (<10 min−1). As shown in Fig. 4C, MBP-depleted Nox2 DH and Nox4 DH(P437H) showed little or no electron transfer activity, similar to the results observed for the MBP-fused forms. Although MBP-depleted Nox4 DH and Nox4 DH mutant (C546L or C547L) catalyzed the NADPH-dependent reduction of cytochrome c or cytochrome b5, respectively, their turnover rates were a little lower than the values observed for MBP-fused Nox4 DH, Nox4 DH(C546L) and Nox4(C547L) proteins. MBP alone showed no activity and the MBP-fused forms of the Nox4 DH domains were used in the present study because of the poor solubility and lower yield of the DH domains when the MBP tag was cleaved and deleted. Little or no Nox4 DH domain-dependent electron transferase activity was observed in the absence of added FAD, indicating loss of FAD during the purification. Fig. 4 View largeDownload slide NADPH-dependent electron transfer activities of wild and mutant MBP-fused Nox DH domain expressed in E. coli—(A) Purified MBP-fusion forms of Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domain (1 μg protein each) were loaded onto 5–20% (w/v) SDS-PAGE gel and stained with Coomassie brilliant blue (a). MBP-depleted form of purified DH proteins (1.5 μg each) were subjected to 5–20% (w/v) SDS-PAGE, followed by protein stain (b). Purified MBP-Nox DH proteins were immunoblotted with anti-body to MBP (c). (B) Pyridine nucreotide-dependent electron transferase activities of purified MBP-fused Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L), Nox4 DH(C547L) and MBP alone were assayed for cytochrome c (white bars) and cytochrome b5 (grey bars) reductase activities. Each reductive activity indicates the average of three independent assays, with the error bars showing the SD. (C) NADPH-dependent electron transfer activities of MBP-depleted Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. The electron transfer assay condition, electron acceptors used and each activity presentation were the same as described in Fig. 4B. Fig. 4 View largeDownload slide NADPH-dependent electron transfer activities of wild and mutant MBP-fused Nox DH domain expressed in E. coli—(A) Purified MBP-fusion forms of Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L) and Nox4 DH(C547L) domain (1 μg protein each) were loaded onto 5–20% (w/v) SDS-PAGE gel and stained with Coomassie brilliant blue (a). MBP-depleted form of purified DH proteins (1.5 μg each) were subjected to 5–20% (w/v) SDS-PAGE, followed by protein stain (b). Purified MBP-Nox DH proteins were immunoblotted with anti-body to MBP (c). (B) Pyridine nucreotide-dependent electron transferase activities of purified MBP-fused Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. Nox2 DH, Nox4 DH, Nox4 DH(P437H), Nox4 DH(C546L), Nox4 DH(C547L) and MBP alone were assayed for cytochrome c (white bars) and cytochrome b5 (grey bars) reductase activities. Each reductive activity indicates the average of three independent assays, with the error bars showing the SD. (C) NADPH-dependent electron transfer activities of MBP-depleted Nox2 DH, Nox4 DH and Nox4 DH mutant proteins. The electron transfer assay condition, electron acceptors used and each activity presentation were the same as described in Fig. 4B. Consistent with previous studies (8, 24), Nox4 DH domain-dependent activities were very low using NADH rather than NADPH as an electron donor. NADPH-dependent cytochrome c or cytochrome b5 reduction by purified Nox4 DH was not largely affected by added SOD (Fig. 5A and B), and is therefore due to a direct electron transfer from the enzyme-bound FAD rather than a superoxide-mediated reaction. The turnover rate of MBP-Nox4 DH(C546L or C547L) also was not remarkably decreased in the presence of added SOD. Addition of DPI and H2O2 strongly decreased NADPH-specific hemeprotein reductase activities of the point mutant Nox4 DH(C546L or C547L) domain as well as wild-type Nox4 DH. In addition, the NADPH concentration-dependent cytochrome c reduction rates of MBP-Nox4 DH and MBP-Nox4 DH mutant (C546L or C547L) were measured (Fig. 5C). The Km value for NADPH was determined to be approximately 17 ± 2 μM in both wild and mutant DH proteins, a value close that previously reported for Nox4 DH (20 ± 5 μM) (8). On the other hand, Vmax values for Nox4 DH(C546L) and Nox4 DH(C547L) mutants were markedly decreased and determined to be 80 ± 5 and 65 ± 5 nmol cytochrome c reduced/min/mg protein, respectively, as compared with Vmax (200 ± 10) of wild-type Nox4 DH. The addition of 50 μM H2O2 considerably decreased the NADPH oxidation rates of MBP-fused Nox4 DH and Nox4 DH mutant (C546L or C547L) proteins in the presence of cytochrome c, whereas the inhibitory effect of H2O2 was significantly lost in the presence of catalase (Fig. 5D). In addition, it was noticed that NADPH oxidative activity of purified Nox4 DH(C547L) mutant was further decreased, about 26% lower than Nox4 DH(C546L) in the absence and presence of 50 μM H2O2 (Fig. 5D). The NADPH oxidation of MBP-Nox4 DH linked with cytochrome b5 reduction was also decreased in the presence of H2O2 (data not shown). Thus, the Nox4 DH domain includes a H2O2-sensing region, and NADPH-specific cytochrome c or cytochrome b5 reductase activity of this domain is clearly regulated by the local H2O2 concentration. Fig. 5 View largeDownload slide Effects of SOD, DPI and H2O2 on NADPH-dependent electron transfer activity by MBP-Nox4 DH or MBP-Nox4 DH mutant—(A) Cytochrome c reduction at 550 nm was measured as described in Figure 4B in the absence and presence of 300 U/ml SOD, 20 μM DPI or 50 μM H2O2. Purified MBP-Nox4 DH (white bars), MBP-Nox4 DH(C546L) (grey bars) or MBP-Nox4 DH(C547L) (black bars) protein was subjected to the assay as indicated. (B) The reduction of cytochrome b5 was quantified by monitoring the absorbance change at 556 nm without and with added SOD, DPI or H2O2 at the indicated concentrations in Fig. 5A. (C) NADPH concentration-dependent cytochrome c reduction of purified MBP-Nox4 DH or MBP-Nox4 DH mutants. Nox4 DH (○), Nox4 DH(C546L, •) and Nox4 DH(C547L, ▵)were assayed for cytochrome c reductase activity, and a double-reciprocal plot of initial velocity versus NADPH concentration on the top was used to determine Km and Vmax values. (D) After purified MBP-Nox4 DH or its mutant form was preincubated with 25 μM FAD and 80 μM cytochrome c in the absence (white bars) or presence of 50 μM H2O2 (grey bars) for 5 min, 50 μM NADPH was added and the rate of NADPH oxidation at 340 nm was assayed at 25°C. NADPH oxidation was also measured in the presence of 50 μM H2O2 just after adding 130 unit/ml of catalase (black bars). (A–D) The data are representative of three separate experiments, with the error bars showing the SD. Fig. 5 View largeDownload slide Effects of SOD, DPI and H2O2 on NADPH-dependent electron transfer activity by MBP-Nox4 DH or MBP-Nox4 DH mutant—(A) Cytochrome c reduction at 550 nm was measured as described in Figure 4B in the absence and presence of 300 U/ml SOD, 20 μM DPI or 50 μM H2O2. Purified MBP-Nox4 DH (white bars), MBP-Nox4 DH(C546L) (grey bars) or MBP-Nox4 DH(C547L) (black bars) protein was subjected to the assay as indicated. (B) The reduction of cytochrome b5 was quantified by monitoring the absorbance change at 556 nm without and with added SOD, DPI or H2O2 at the indicated concentrations in Fig. 5A. (C) NADPH concentration-dependent cytochrome c reduction of purified MBP-Nox4 DH or MBP-Nox4 DH mutants. Nox4 DH (○), Nox4 DH(C546L, •) and Nox4 DH(C547L, ▵)were assayed for cytochrome c reductase activity, and a double-reciprocal plot of initial velocity versus NADPH concentration on the top was used to determine Km and Vmax values. (D) After purified MBP-Nox4 DH or its mutant form was preincubated with 25 μM FAD and 80 μM cytochrome c in the absence (white bars) or presence of 50 μM H2O2 (grey bars) for 5 min, 50 μM NADPH was added and the rate of NADPH oxidation at 340 nm was assayed at 25°C. NADPH oxidation was also measured in the presence of 50 μM H2O2 just after adding 130 unit/ml of catalase (black bars). (A–D) The data are representative of three separate experiments, with the error bars showing the SD. Discussion Purified Nox4 DH-dependent reduction of heme protein such as cytochrome c or cytochrome b5 supports a model for the endogeneous NADPH-to-FAD-to-heme bL-to-heme bH electron transfer that occurs in the holoenzyme of Nox4 flavocytochrome. In this model, we were able to indicate the correspondence in H2O2 effect on intact Nox4 turnover during the NADPH-dependent ROS formation (Fig. 6A) and NADPH-dependent electron transfer rates to artificial hemeproteins (Fig. 6B), suggesting that cytosolic H2O2 regulates the rate-limiting FAD reduction in holo-Nox4 enzyme. Thus, the present data provide a reaction system in which H2O2 concentration-dependent reversible change of Nox4 DH domain structure appears to be co-regulated by peroxidases such as TRx, GRx and catalase (Fig. 6C). Fig. 6 View largeDownload slide Hydrogen peroxide-dependent regulation of constitutive active Nox4 enzyme—(A) Hydride ion transfer from NADPH to FAD is assumed to be the rate-determining step in Nox4 flavocytochrome, which is regulated by cytosolic H2O2 concentration. After the reduction of FAD by NADPH, the DH domain/TM-heme domain interaction may help orient and localize the FAD in close proximity to heme bL, thus allowing rapid electron transfer. (B) The electron transfer rate from purified Nox4 DH toward artificial heme proteins is affected in the presence of physiological range of H2O2. (C) Nox4 activity is affected by H2O2 concentration-dependent Cys-SH residue(s) oxidation of DH domain, which is adjusted by cytosolic peroxidases such as thioredoxin (TRx), glutaredoxin (GRx), or catalase. Fig. 6 View largeDownload slide Hydrogen peroxide-dependent regulation of constitutive active Nox4 enzyme—(A) Hydride ion transfer from NADPH to FAD is assumed to be the rate-determining step in Nox4 flavocytochrome, which is regulated by cytosolic H2O2 concentration. After the reduction of FAD by NADPH, the DH domain/TM-heme domain interaction may help orient and localize the FAD in close proximity to heme bL, thus allowing rapid electron transfer. (B) The electron transfer rate from purified Nox4 DH toward artificial heme proteins is affected in the presence of physiological range of H2O2. (C) Nox4 activity is affected by H2O2 concentration-dependent Cys-SH residue(s) oxidation of DH domain, which is adjusted by cytosolic peroxidases such as thioredoxin (TRx), glutaredoxin (GRx), or catalase. Although Nox4 expression is greatest in the kidney (23, 31, 32), it is widely expressed in many other cell types (32–34) and hence may have a cellular function that is more general than that of other Nox enzymes with more restricted tissue expression. We found that transfection of His6-tagged Nox4 cDNA to HEK293 cells induced further synthesis of p22phox subunit and increased Nox4-p22phox complex was observed in nuclear membrane fraction (Fig. 2A), which agrees with previous reports that stabilized p22phox expression responds to the level of constitutively active Nox4 formation and its intracellular localization (35). Thus, the Nox4 flavocytochrome-p22phox interaction produces a more stable conformation of the Nox4 protein localized to the nuclear membranes of HEK293 cells for the catalytic function to produce ROS. A recent study reported the major product from Nox4 is H2O2, and its activity is regulated by both its expression level and oxygen availability, functioning as an oxygen sensor (9). In fact, Nox4 generated H2O2 approximately in direct proportion to oxygen at concentrations below about 10%, making it a sensitive reporter of tissue oxygenation level. That Nox4 shows a high Km value for oxygen and generates mostly hydrogen peroxide indicates that it responds to the output of H2O2 rather than to external signals via intermediate signalling mechanisms, such as changes in cellular Ca2+ concentration or phosphorylation of regulatory subunits. Hydrogen peroxide shows relative specificity toward low pKa cysteine residues allowing its use as a signalling molecule to regulate enzyme activity (36, 37). We suggest that H2O2 participates in signalling ROS to regulate Nox4 activity in adaptive response to oxidative stress or hypoxic condition. Several studies indicate that specific location of Cys residues in proteins is associated with subcellular function (38, 39), suggesting a basic consideration of framework for differential protein oxidation due to oxidative stresss and toxicity linked with redox response and function. The reactive thiol side chain of Cys residues can function as a sensor or switch, changing between the reduced and oxidized state in response to fluctuations in ROS. Depending on the local ROS concentration, hydrogen peroxide, superoxide or NO can react with Cys residue to form reversible modifications (S-sulfenylation, disulfide bond formation, or S-nitrosylation). The introduction of new disulfide formation has the potential significantly to alter protein conformation and affect bio-function. Many proteins with reactive cysteines are involved in controlling thiol-disulfide exchange reactions to regulate enzyme activity and maintain cellular redox balance (40–42). In the steady state, thiols undergo oxidation due to increased oxidant such as H2O2 produced by Nox enzymes, suggesting that reactive peptidyl cysteine oxidation is associated with functional signal networks (43). Conceivably, in the nuclei, reversible structural change of redox-sensitive protein has a critical role for the activation or inactivation of transcriptional factors (44, 45). Hydrogen peroxide can cross membranes and is relatively stable. Although hydrogen peroxide has been intrinsically associated with oxidative stress, the physiological role of this mild oxidant may be as a redox signal messenger, and its production is likely controlled by the H2O2-sensitive Nox4 enzyme (46, 47). In fact, the redox sensitive thiol side chain oxidation of Cys residues in Nox4 DH domain appears to affect the Nox4 activity. Since NADPH-dependent electron transfer rate of the DH domain to heme protein such as cytochrome c or cytochrome b5 was remarkably affected by H2O2, the reversible oxidative modification of conserved and reactive Cys residues by H2O2 are assumed to be responsive to Nox4 activity. The NADPH oxidation rate of either wild-type or mutant Nox4 expressed in the membranes of NEF was clearly decreased with increasing H2O2 concentration (Fig. 3C and D). On the other hand, NADPH-dependent reduction of the artificial electron acceptor by either Nox4(C546L) or Nox4(C547L) mutant DH was significantly lower rate and less sensitive to H2O2 (Fig. 5A and B). In accord with these results of the mutant Nox4 DH, the decrease of H2O2-producing activity linked with NADPH oxidation rate of Nox4(C546L or C547L) flavocytochrome was also implied in NEF (Figs 2C and 3B). Our results, therefore, suggest that H2O2 production rate by constitutively active Nox4 should be controlled in intracellular H2O2 concentration. The increased H2O2 production with time in the nuclei caused by Nox4 up-regulation may induce oxidative nuclear damage leading to dysfunction and apoptotic cell death; therefore, the activity of Nox4 should be constantly auto-regulated by its DH flavoprotein domain. Since H2O2 reacts with Fe2+ and produces hydroxy radical (.OH) which enhances the risk of intracellular damages, it is important to regulate the H2O2 generating activity of Nox4 enzyme. On the other hand, anti-oxidant enzymes such as catalase, thioredoxin and glutaredoxin are considered to preferentially reduce sulfenic acid or disulfide bridge formation of Cys residues in DH domain of Nox4 to keep thermodynamic stability of the enzyme activity (Fig. 6C). In view of the three-dimensional protein model on the basis of its primary structure (48), the Nox4 C-terminal DH domain was shown to be composed of two units, a β-fold capped by a single helix to bind FAD, followed by a Rossmann-fold that binds NADPH (Fig. 7). In this model, Cys546 or/and Cys547 are suggested to be part of the NADPH binding site based on homology to ferredoxin-NADP+ reductase, cytochrome P-450 reductase and nitric oxide synthase (49, 50). When the C-terminal conserved amino acid sequences of NADPH binding sites are compared among human Nox family members (34), the Cys546-Cys547 sequence is located specifically in only the Nox4 DH domain (Table I). Thus, the hydrogen peroxide-dependent sulfenic acid or disulfide formation of these Cys residues may induce a limited reversible change of the DH domain conformation affecting the ROS producing turnover of constitutively active Nox4 enzyme. Table I. C-terminal conserved putative NADPH-binding amino acid sequences of human Nox isoform— Underline indicates Nox4 DH specific conserved Cys546/Cys547 residues that are predicted to control NADPH DH activity in response to intracellular H2O2 concentration Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Table I. C-terminal conserved putative NADPH-binding amino acid sequences of human Nox isoform— Underline indicates Nox4 DH specific conserved Cys546/Cys547 residues that are predicted to control NADPH DH activity in response to intracellular H2O2 concentration Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Nox1—526 VGVFLCGP533—  Nox2—533 IGVFLCGP540—  Nox3—530 IGVFFCGP537—  Nox4—542 VGVFCCGP549—  Nox5—535 VQVFFCGS542—  Duox1–1515IGVFSCGP1522—  Duox2–1512IGVFSCGP1519—  Fig. 7 View largeDownload slide Three-dimensional model of Nox4—A three-dimensional model is given for Nox4 based on its primary structure (48). The Nox4 C-terminal DH domain is composed of two units, a β-fold capped by a single helix that binds FAD, and a Rossman-fold that binds NADPH. Cys546 and Cys547 are localized in NADPH binding site. Presumably, the hydrogen peroxide-dependent sulfenic acid or disulfide formation between these two Cys residues may induce a limited reversible change in the conformation of the DH domain that affects turnover of the ROS-producing Nox4 enzyme. Fig. 7 View largeDownload slide Three-dimensional model of Nox4—A three-dimensional model is given for Nox4 based on its primary structure (48). The Nox4 C-terminal DH domain is composed of two units, a β-fold capped by a single helix that binds FAD, and a Rossman-fold that binds NADPH. Cys546 and Cys547 are localized in NADPH binding site. Presumably, the hydrogen peroxide-dependent sulfenic acid or disulfide formation between these two Cys residues may induce a limited reversible change in the conformation of the DH domain that affects turnover of the ROS-producing Nox4 enzyme. Acknowledgement The authors express appreciate to Dr Toshitsugu Yubisui (Department of Biochemistry, Okayama University of Science, Okayama, Japan) for the gift of soluble purified cytochrome b5 protein. Funding The present study was fully supported by the research grant of applied science to educate and direct graduate in both Chubu University and Fujita Health University. No additional financial support is not added to this present research focused on the enzyme function. Conflict of Interest None declared. References 1 D’Autréaux B., Toledano M.B. ( 2007) ROS as signaling molecules: mechanisms that generate specificity in ROS homeostasis. Nat. Rev. Mol. Cell Biol . 8, 813– 824 Google Scholar CrossRef Search ADS PubMed  2 Finkel T. ( 2011) Signal transduction by reactive oxygen species. J. Cell Biol.  194, 7– 15 Google Scholar CrossRef Search ADS PubMed  3 Turpaev K.T. ( 2002) Reactive oxygen species and regulation of gene expression. Biochemistry  67, 281– 292 Google Scholar PubMed  4 Bae Y.S., Oh H., Rhee S.G., Yoo Y.D. ( 2011) Regulation of reactive oxygen species generation in cell signaling. Mol. Cells  32, 491– 509 Google Scholar CrossRef Search ADS PubMed  5 Bánfi B., Molnár G., Maturana A., Steger K., Hegedûs B., Demaurex N., Krause K.-H. ( 2001) ACa2+-activated NADPH oxidase in testis, spleen, and lymph nodes. J. Biol. Chem . 276, 37594– 37601 Google Scholar CrossRef Search ADS PubMed  6 Bánfi B., Tirone F., Durussel I., Knisz J., Moskwa P., Molnár G.Z., Krause K.-H., Cox J.A. ( 2004) Mechanism of Ca2+ activation of the NADPH oxidase 5 (NOX5). J. Biol. Chem . 279, 18583– 18591 Google Scholar CrossRef Search ADS PubMed  7 Bánfi B., Malgrange B., Knisz J., Steger K., Dubois-Dauphin M., Krause K.-H. ( 2004) NOX3, a superoxide-generating NADPH oxidase of the inner ear. J. Biol. Chem . 279, 46065– 46072 Google Scholar CrossRef Search ADS PubMed  8 Nisimoto Y., Jackson M.H., Ogawa H., Kawahara T., Lambeth J.D. ( 2010) Constitutive NADPH dependent electron transferase activity of the Nox4 dehydrogenase domain. Biochemistry  49, 2433– 2442 Google Scholar CrossRef Search ADS PubMed  9 Nisimoto Y., Diebold A.B., Gomes C.D., Lambeth J.D. ( 2014) Nox4: a hydrogen peroxide-generating oxygen sensor. Biochemistry  53, 5111– 5120 Google Scholar CrossRef Search ADS PubMed  10 Ambasta R.K., Kumar P., Griendling K.K., Schmidt H.H., Busse R., Brandes R.P. ( 2004) Direct interaction of the novel Nox proteins with p22phox is required for the formation of a functionally active NADPH oxidase. J. Biol. Chem . 279, 45935– 45941 Google Scholar CrossRef Search ADS PubMed  11 Parkos C.A., Dinauer M.C., Walker L.E., Allen R.A., Jesaitis A.J., Orkin S.H. ( 1988) Primary structure and unique expression of the 22-kilodalton light chain of human neutrophil cytochrome b. Proc. Natl. Acad. Sci. USA  85, 3319– 3323 Google Scholar CrossRef Search ADS   12 Zhu Y., Marchal C.C., Cashbon A.J., Stull N., von Lohneysen K., Kraus U.C., Jesaitis A.J., McCmick S., Nauseef W.M., Dinauer M.C. ( 2006) Deletion of p22phox subunit of flavocytochrome b558: identification of regions critical for gp91phox maturation and NADPH oxidase activity. J. Biol.Chem . 281, 30336– 30346 Google Scholar CrossRef Search ADS PubMed  13 Yu L., Zhen L., Dinauer M.C. ( 1997) Biosynthesis of the phagocyte NADPH oxidase cytochrome b558: role of heme incorporation and heterodimer formation in maturation and stability of gp91phox and p22phox subunits. J. Biol. Chem . 272, 27286– 27294 14 Nakano Y., Banfi B., Jesaitis A.J., Dinauer M.C., Allen L.-A.H., Nauseef W.M. ( 2007) Critical roles for p22phox in the structural maturation and subcellular targeting of Nox3. Biochem. J . 403, 97– 108 Google Scholar CrossRef Search ADS PubMed  15 Martyn K.D., Frederick L.M., von Loehneysen K., Dinauer M.C., Knaus U.G. ( 2006) Functional analysis of Nox4 reveals unique characteristics compared to other NADPH oxidase. Cell Signal  18, 69– 82 Google Scholar CrossRef Search ADS PubMed  16 Serrander L., Cartierr L., Bendard K., Banfi B., Lardy B., Plastre O., Sienkiewicz A., Forro L., Schlegel W., Krause K.-H. ( 2007) NOX4 activity is determined by mRNA levels and reveals a unique pattern of ROS generation. Biochem. J . 406, 105– 114 Google Scholar CrossRef Search ADS PubMed  17 Krause K.-H. ( 2004) Tissue distribution and putative physiological function of NOX family NADPH oxidase. Jpn. J. Infect. Dis . 57, S28– S29 Google Scholar PubMed  18 Lambeth J.D., Kawahara T., Diebold B. ( 2007) Regulation of Nox and Duox enzymatic activity and expression. Free Radical Biol. Med . 43, 319– 331 Google Scholar CrossRef Search ADS   19 Ago T., Kuroda J., Pain J., Fu C., Li H., Sadoshima J. ( 2010) Upregulation of Nox4 by hypertrophic stimuli promotes apoptosis and mitochondrial dysfunction in cardiac myocytes. Circ. Res . 106, 1– 12 Google Scholar CrossRef Search ADS   20 Takac I., Schröder K., Zhang L., Lardy B., Anilkuman N., Lambeth J.D., Shah A.M., Morel F., Brandes R.P. ( 2011) The E-loop is involved in hydrogen peroxide formation by the NADPH oxidase Nox4. J. Biol. Chem . 286, 13304– 13313 Google Scholar CrossRef Search ADS PubMed  21 Van Buul J.D., Fernandez-Borja M., Anthony E.C., Hordijk P.L. ( 2005) Expression and localization of Nox2 and Nox4 in primary human endothelial cells. Antioxidant Redox Signal . 7, 308– 317 Google Scholar CrossRef Search ADS   22 Hilenski L.L., Clempus R.E., Quinn M.T., Lambeth J.D., Griendling K.K. ( 2004) Distinct subcellular localizations of Nox1 and Nox4 in vascular smooth muscle cells. Arterioscler. Thromb. Vasc. Biol . 24, 677– 683 Google Scholar CrossRef Search ADS PubMed  23 Geiszt M., Kopp J.B., Varnai P., Leto T.L. ( 2000) Identification of renox, an NAD(P)H oxidase in kidney. Proc. Natl. Acad. Sci. USA  97, 8010– 8014 Google Scholar CrossRef Search ADS   24 Jackson H.M., Kawahara T., Nisimoto Y., Smith S.M.E., Lambeth J.D. ( 2010) Nox4 B-loop creates an interface between the transmembrane and dehydrogenase domains. J. Biol. Chem . 285, 10281– 10290 Google Scholar CrossRef Search ADS PubMed  25 Nisimoto Y., Ogawa H., Miyano K., Tamura M. ( 2004) Activation of the flavoprotein domain of gp91phox upon interaction with N-terminal p67phox (1- 210) and the Rac complex. Biochemistry  43, 9567– 9575 Google Scholar CrossRef Search ADS PubMed  26 Lutter R., van Schaik M.L.J., van Zwieten R., Wever R., Roos D., Hamers M.N. ( 1985) Purification and partial characterization of the b-type cytochrome from human polymorphonuclear Leukocytes. J. Biol. Chem . 260, 2237– 2244 Google Scholar PubMed  27 Vermilion J.L., Ballou D.P., Massey V., Coon M.J. ( 1981) Separate roles for FMN and FAD in catalysis by liver microsomal NADPH-cytochrome P-450 reductase. J. Biol. Chem.  256, 266– 277 Google Scholar PubMed  28 Zhao B., Summers F.A., Mason R.P. ( 2012) Photooxidation of Amplex Red to resorufin: implications of exposing the Amplex Red assay to light. Free Radic. Biol. Med . 53, 1080– 1087 Google Scholar CrossRef Search ADS PubMed  29 Rodrigues J.V., Gomes C.M. ( 2010) Enhanced superoxide and hydrogen peroxide detection in biological assays. Free Radical Biol. Med . 49, 61– 66 Google Scholar CrossRef Search ADS   30 Kimura S., Nishida H., Iyanagi T. ( 2001) Effects of flavin-binding motif amino acid mutations in the NADH-cytochrme b5 reductase catalytic domain on protein stability and catalysis. J. Biochem . 130, 481– 490 Google Scholar CrossRef Search ADS PubMed  31 Cheng G., Cao Z., Xu X., Meir E.G.V., Lambeth J.D. ( 2001) Homologs of gp91phox: cloning and tissue expression of Nox3, Nox4, and Nox5. Gene  269, 131– 140 Google Scholar CrossRef Search ADS PubMed  32 Shiose A., Kuroda J., Tsuruya K., Hirai M., Hirakata H., Naito S., Hattori M., Sakaki Y., Sumimoto H. ( 2001) A novel superoxide-producing NAD(P)H oxidase in kidney. J. Biol. Chem.  276, 1417– 1423 Google Scholar CrossRef Search ADS PubMed  33 Lambeth J.D. ( 2004) NOX enzymes and the biology of reactive oxygen. Nat. Rev. Immunol . 4, 181 Google Scholar CrossRef Search ADS PubMed  34 Lambeth J.D., Cheng G., Arnold R.S., Edens W.E. ( 2000) Novel homologs of gp91phox. Trends Biochem. Sci . 25, 459– 461 Google Scholar CrossRef Search ADS PubMed  35 von Löhneysen K., Noack D., Jesaitis A.J., Dinauer M.C., Knaus U.G. ( 2008) Mutational analysis reveals distinct features of the Nox4-p22phox complex. J. Biol. Chem.  283, 35273– 35282 Google Scholar CrossRef Search ADS PubMed  36 Denu J.M., Tanner K.G. ( 1998) Specific and reversible inactivation of tyrosine phosphatase by hydrogen peroxide: evidence for a sulfenic acid intermediate and implications for redox regulation. Biochemistry  37, 5633– 5642 Google Scholar CrossRef Search ADS PubMed  37 Barrett W.C., DeGnore J.P., König S., Fales H.M., Keng Y.-F., Zhang Y., Yim M.B., Chock P.B. ( 1999) Regulation of PTP1B via glutathionylation of the active site cysteine. Biochemistry  38, 6699– 6705 Google Scholar CrossRef Search ADS PubMed  38 Ziegler D.M. ( 1985) Role of reversible oxidation-reduction of enzyme thiols-disulfides in metabolic regulation. Annu. Rev. Biochem.  54, 305– 329 Google Scholar CrossRef Search ADS PubMed  39 Gilbert H.F. ( 1995) Thiol/disulfide exchange equilibria and disulfide bond stability. Methods Enzymol . 251, 8– 28 Google Scholar CrossRef Search ADS PubMed  40 Jones D.P., Go Y.M., Anderson C.L., Ziegler T.R., Kinkade J.M., Kirlin W.G. ( 2004) Cysteine/cystine couple is a newly recognized node in the circuitry for biologic redox signaling and Control. Faseb J . 18, 1246– 1248 Google Scholar CrossRef Search ADS PubMed  41 Netto L.E.S., Oliveira M.A., Monteiro G., Demasi A.P.D., Cussiol J.R.R., Discola K.F., Demasi M., Silva G.M., Alves S.V., Faria V.G., Horta B.B. ( 2007) Reactive cysteine in proteins: protein folding, antioxidant defense, redox signaling and more. Comparative Biochem. Physiol. Part C  146, 180– 193 42 Go Y.M., Jones D.P. ( 2010) Redox control systems in the nucleus: mechanisms and functions. Antioxid. Redox Signal . 13, 489– 509 Google Scholar CrossRef Search ADS PubMed  43 Miki H., Funato Y. ( 2012) Regulation of intracellular signaling through cysteine oxidation by reactive oxygen species. J. Biochem . 151, 255– 261 Google Scholar CrossRef Search ADS PubMed  44 Haddad J.J. ( 2002) Science review: redox and oxygen-sensitive transcription factors in the regulation of oxidant-mediated lung injury: role for nuclear factor-kB. Crit. Care  6, 481– 490 Google Scholar CrossRef Search ADS PubMed  45 Merchant A.A., Singh A., Matsui W., Biswal S. ( 2011) The redox-sensitive transcription factor Nrf2 regulates murine hematopoietic stem cell survival independently of ROS levels. Blood  118, 6572– 6579 Google Scholar CrossRef Search ADS PubMed  46 Rhee S.G. ( 1999) Redox signaling: hydrogen peroxide as intracellular messenger. Exp. Mol. Med . 31, 53– 59 Google Scholar CrossRef Search ADS PubMed  47 Stone J.R., Yang S. ( 2006) Hydrogen peroxide: a signaling messenger. Antioxid. Redox Signal  8, 243– 270 Google Scholar CrossRef Search ADS PubMed  48 Jackson H.M. ( 2010) Structure and Functional Analysis of NADPH Oxidase 4. Ph. D. thesis. Emory University, ATL, pp. 1– 210 49 Segal A.W., West I., Wientjes F.B., Nugent J.H.A., Chavan A.J., Haley B., Garcia R.C., Rosen H., Scrace G. ( 1992) Cytochrome b-245 is a flavocytochrome containing FAD and the NADPH-binding site of the microbicidal oxidase of phagocytes. Biochem. J.  284, 781– 788 Google Scholar CrossRef Search ADS PubMed  50 Taylor W.R., Jones D.T., Segal A.W. ( 1993) A structural model for the nucleotide binding domains of the flavocytochrome b-245 beta-chain. Protein Sci . 2, 1675– 1685 Google Scholar CrossRef Search ADS PubMed  Abbreviations Abbreviations DH dehydrogenase DPI diphenylene iodonium Duox dual oxidase GRx glutaredoxin HEK human embryonic kidney HRP horseradish peroxidase MBP maltose binding protein NEF nucleus-enriched fraction Nox NADPH oxidase PMSF phenylmethanesulfonyl fluoride phox phagocite oxidase ROS reactive oxygen species SOD superoxide dismutase TM transmembrane TRx thioredoxin © The Author(s) 2018. Published by Oxford University Press on behalf of the Japanese Biochemical Society. All rights reserved This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/about_us/legal/notices)

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The Journal of BiochemistryOxford University Press

Published: Jan 21, 2018

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