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Inactivation of Human Arylamine N‐Acetyltransferase 1 by Hydrogen Peroxide and Peroxynitrite
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Wild, D., Feser, W., Michel, S., Lord, H. L., and Josephy, P. D. (1995). Metabolic activation of heterocyclic aromatic amines catalyzed by human arylamine N‐acetyltransferase isozymes (NAT1 and NAT2) expressed in Salmonella typhimurium. Carcinogenesis 16, 643–648. Xenarios, I., Salwinski, L., Duan, X. J., Higney, P., Kim, S. M., and Eisenberg, D. (2002). DIP, the Database of Interacting Proteins: A research tool for studying cellular networks of protein interactions. Nucleic Acids Res. 30, 303–305. Zhou, H., Josephy, P. D., Kim, D., and Guengerich, F. P. (2004). Functional characterization of four allelic variants of human cytochrome P450 1A2. Arch. Biochem. Biophys. 422, 23–30.
[12] Inactivation of Human Arylamine N‐Acetyltransferase 1 by Hydrogen Peroxide and Peroxynitrite By JEAN‐MARIE DUPRET, JULIEN DAIROU, NOUREDDINE ATMANE , and FERNANDO RODRIGUES‐LIMA Abstract
Arylamine N‐acetyltransferases (NAT) are xenobiotic‐metabolizing enzymes responsible for the acetylation of many arylamine and heterocyclic amines. They therefore play an important role in the detoxification and activation of numerous drugs and carcinogens. Two closely related isoforms (NAT1 and NAT2) have been described in humans. NAT2 is present mainly in the liver and intestine, whereas NAT1 is found in a wide range of tissues. Interindividual variations in NAT genes have been shown to be a potential source of pharmacological and/or pathological susceptibility. Evidence now shows that redox conditions may also contribute to overall NAT activity. This chapter summarizes current knowledge on human NAT1 regulation by reactive oxygen and nitrogen species. Introduction
Arylamine N‐acetyltransferases (NATs) are xenobiotic‐metabolizing enzymes (XME) that catalyze the transfer of an acetyl moiety from acetyl‐CoA to the nitrogen atom of primary arylamines and hydrazines. They are also responsible for the O‐acetylation of N‐hydroxyarylamines (Pompeo et al., 2002). NATs therefore participate in the detoxification and/or activation of a variety of drugs and carcinogens (Badawi et al., 1995; Hein, 2002). NATs have been identified in several species, ranging from bacteria to mammals (Delome´nie et al., 2001; Payton et al., 2001; Rodrigues‐Lima and METHODS IN ENZYMOLOGY, VOL. 400 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)00012-1
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Dupret, 2002b). Two functional isoforms of NAT (NAT1 and NAT2) have been described in humans (Matas et al., 1997). NAT1 is found in a wide range of tissues and organs (Rodrigues‐Lima et al., 2003). NAT2 is present mainly in the liver and intestine (Hickman et al., 1998). However, it is likely that NAT2 expression is more widespread (Dairou et al., 2005; Dupret and Rodrigues‐Lima, 2005). Genetically determined interindividual variation in NAT2 content and activity is the basis of the well‐known isoniazid acetylation polymorphism. Polymorphism associated with the NAT1 isoform may also be a source of pharmacological susceptibility (Hein et al., 2000). To date, at least 26 NAT1 alleles and 36 NAT2 alleles have been identified, resulting from numerous single‐nucleotide polymorphisms (Dupret et al., 2004). A detailed description of NAT alleles is available at http://www.louisville.edu/medschool/ pharmacology/NAT.html. Although the relationships between polymorphic substitutions and the activity of variant enzymes have been investigated thoroughly (Fretland et al., 2001a,b), further studies are required to determine the effects of nucleotide variations outside the NAT coding regions and to characterize the enzyme activity of allelic variants more fully. Studies have shown that other mechanisms, such as substrate‐dependent downregulation (Butcher et al., 2000, 2004), the existence of splice variants (Husain et al., 2004), and posttranslational inhibition of NAT activity (Atmane et al., 2003; Dairou et al., 2005), may also contribute to the overall activity of NATs. This chapter summarizes current knowledge on NAT inhibition by reactive oxygen or nitrogen species, consistent with the fundamental role in NAT‐mediated catalysis of a conserved active‐site residue. NAT‐Mediated Catalysis
Results of early steady‐state kinetic studies were consistent with a simple two‐step substituted enzyme (‘‘ping‐pong’’) kinetic mechanism for the NAT reaction (Riddle and Jencks, 1971). Subsequent site‐directed mutagenesis and functional studies have shown that an acetylcysteinyl enzyme is involved in the catalytic process (Dupret and Grant, 1992). Finally, the crystallographic determination of the structure of NATs from prokaryotic species (Salmonella typhimurium, Mycobacterium smegmatis, Pseudomonas aeruginosa, Mesorhizobium loti) (Holton et al., 2005; Sandy et al., 2002; Sinclair et al., 2000; Westwood et al., 2005) and the construction of homology models of human NAT1 and NAT2 (Rodrigues‐Lima and Dupret, 2002a; Rodrigues‐Lima et al., 2001) have revealed structural similarities with cysteine proteases. These studies revealed the existence of a protease‐like catalytic triad (residues Cys 68‐His 107‐Asp 122 in humans) (Fig. 1). The strict conservation of these structural features suggests that
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FIG. 1. Ribbon‐and‐stick representation of the human NAT1 catalytic triad. This figure was created with Deep‐View 3.7b2 (Guex and Peitsch, 1997) using the homology model of the N‐terminal domain of human NAT1 (Rodrigues‐Lima et al., 2001). The hydrogen bonds among cysteine, histidine, and aspartate residues are shown as dotted lines.
NATs have adapted a catalytic mechanism commonly found in cysteine proteases for use in the acetyl‐transfer reaction. Although the exact catalytic mechanism of NAT enzymes is not fully understood, it has been shown to depend on the formation of a thiolate‐ imidazolium ion pair between the triad cysteine and histidine (Wang et al., 2004). The use of chemical modification procedures to probe the reactivity of the active‐site cysteine revealed that it undergoes alkylation upon NAT1 labeling with N‐arylbromoacetamido reagents (Guo et al., 2003). This type of covalent modification can also occur upon NAT1‐catalyzed bioactivation of N‐hydroxy‐2‐acetylaminofluorene and subsequent sulfinamide adduct formation (Guo et al., 2004). In addition to its importance as the primary site of covalent modification in NAT1, Cys 68 has been shown to be the molecular target of reactive oxygen or nitrogen species (Atmane et al., 2003; Dairou et al., 2003, 2004). Oxidative Inhibition of NATs
Several enzymes with a reactive catalytic cysteine residue have been shown to be regulated by the reactive oxygen or nitrogen species (ROS and RNS, respectively) generated during oxidative stress (Halliwell and Gutteridge, 1999). Hydrogen peroxide (H2O2), one of the major cellular
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oxidants, inactivates phosphatases both in vitro and in vivo (Caselli et al., 1998; Lee et al., 1998, 2002). Generally, in vivo and in vitro, H2O2 can oxidize the cysteine residue to give cysteine sulfenic acid (cys‐SOH) or disulfide, which can be reduced back to cysteine by cellular reductants (Halliwell and Gutteridge, 1999). S‐nitrosothiols (RSNO) are reactive nitrogen species known to inactivate cysteine‐containing enzymes, such as papain (Xian et al., 2000a), phosphatases (Xian et al., 2000b), and transglutaminases (Bernassola et al., 1999), either through S‐nitrosylation (formation of an S‐NO) or through formation of a mixed disulfide bond. Both modifications can be reversed by reducing conditions. Peroxynitrite (ONOO) also affects protein function by modifying essential reactive thiols or tyrosine residues (Groves, 1999; Radi et al., 1991). It irreversibly inactivates several enzymes, including creatine kinase (Konorev et al., 1998), caspases (Mohr et al., 1997), and phosphatases (Takakura et al., 1999). XMEs such as cytochrome P450 (Lin et al., 2003) and glutathione S‐transferases (Wong et al., 2001) are also irreversibly inactivated by peroxynitrite. We assessed the ability of reactive oxygen and nitrogen species to inactivate human NAT1. We also investigated the possible reactivation of oxidized NAT1 by reducing agents. Our observations led us to propose that H2O2 and RSNO reversibly inhibit human NAT1, whereas peroxynitrite irreversibly inhibits the enzyme (Rodrigues‐Lima and Dupret, 2004) (Fig. 2).
FIG. 2. Schematic representation of the regulation of human NAT1 by oxidation. The catalytic cysteine residue of the reduced active form of NAT1 is present predominantly as a thiolate anion. This enhances its nucleophilic properties but renders NAT1 susceptible to oxidation. Low levels of H2O2 lead to the reversible oxidation of the catalytic cysteine residue to a sulfenic acid (SOH) with concomitant reversible inactivation of NAT1 activity. When NAT1 is oxidized by S‐nitrosothiols (RSNO), a mixed disulfide (NAT1‐S‐SR) is formed at the catalytic cysteine residue with reversible inactivation of NAT1. Oxidation by low concentrations of peroxynitrite leads to the irreversible inactivation of NAT1 via oxidation of the catalytic cysteine to a sulfinic (SO2H) or sulfonic acid (SO3H).
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Here, we describe in vitro and ex vivo approaches that allow the study of the effect of ROS or RNS on the activity of human NAT1. These reactive species are produced in response to a wide variety of stimuli, such as exposure to chemicals, hormones, and ultraviolet, and following exposure to deleterious physiological/physiopathological conditions, such as apoptosis and inflammation (Groves, 1999; Halliwell and Gutteridge, 1999; Liu and Stamler, 1999; Tannickal and Fanburg, 2000). Excessive production of ROS and RNS is known to participate in the pathological processes of several diseases such as cancer and cataract (Halliwell and Gutteridge, 1999). In addition, these oxidants may affect XME‐dependent biotransformation (Boelsterli, 2003; Pagano, 2002). Thus, the protocols described herein may be used to study the sensitivity of NATs toward oxidative stress induced by various types of stimulus.
General Methods
Expression and Purification of Recombinant Human NAT1 To study the effect of oxidants on recombinant human NAT1, the enzyme is expressed (PET28A vector from Novagen) as a polyhistidine‐ tagged fusion protein (His‐NAT1) in Escherichia coli BL21(DE3) grown for 4 h at 37 in the presence of 0.5 mM isopropyl‐1‐thio‐ ‐D‐galactopyranoside. To purify recombinant NAT1, lysates are loaded onto nickel‐ agarose affinity chromatography columns. Purified His‐NAT1 is reduced by treating with 10 mM dithiothreitol (DTT) (final concentration) for 10 min at 4 prior to dialysis against 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA. SDS–PAGE analysis is carried out at each stage of purification, and protein concentrations are determined using a standard Bradford assay. Cells Expressing NAT1 The effects of oxidants on endogenous cellular NAT1 have been studied using MCF7 (human mammary carcinoma) cells, which are known to express NAT1 (Adam et al., 2003). Other cells have also been used to study the effects of oxidants on endogenous NAT1 (Dairou et al., 2005). Given that the NAT1 gene appears to be expressed ubiquitously, it is likely that most cell types could be used. NAT1 Assays NAT1 activity can be detected easily using spectrophotometric assays. HPLC assays may also be used (Dupret and Grant, 1992). Only
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spectrophotometric assays are described here. These assays can be done easily in the wells of microtiter plates. First Method. NAT1 enzyme activity is determined spectrophotometrically (410 nm) using p‐nitrophenylacetate (PNPA) as the acetyl donor and a NAT1‐specific arylamine substrate such as p‐aminosalicylic acid (PAS) (Musthaq et al., 2002). Briefly, oxidized and unoxidized forms of NAT1 (10–50 l) are assayed in a reaction mixture containing 500 M PAS (final concentration) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA. Reactions are started by adding 125 M PNPA (final concentration). Total reaction volume is 1 ml and final enzyme concentration is usually 10–20 nM. After 10 min at 37 , reactions are quenched by SDS (1% final concentration). p‐Nitrophenol, generated through hydrolysis of PNPA by NAT1 in presence of PAS, is quantified by measuring absorbance at 410 nm with an ELISA plate analyzer. One unit of p‐nitrophenol is defined as the amount of enzyme that gives an absorbance at 410 nm of 0.5 per 10 min/ml. Controls are included without enzyme, PNPA, or PAS. All enzymatic reactions are performed in quadruplicate under conditions where the initial rates are linear. Second Method. The rate of hydrolysis of acetyl‐CoA (first substrate) by NAT1 in the presence of the NAT1‐specific arylamine substrate (PAS) can be determined by detecting the free CoA thiol with 5,50 ‐dithio‐bis 2‐ nitrobenzoic acid (DTNB or Ellman’s reagent) as described previously by Brooke et al. (2003). PAS (500 M final) and samples containing purified NAT1 (0.25 g enzyme) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA buffer are mixed and preincubated (37 , 5 min) in a 96‐well ELISA plate. Acetyl‐ CoA is added to a final concentration of 400 M to start the reaction in a total volume of 100 l. The reaction is quenched with 25 l of 5 mM DTNB in guanidine hydrochloride solution (6.4 M guanidine‐HCl, 0.1 M Tris‐HCl, pH 7.3). The absorbance at 405 nm is measured using an ELISA plate analyzer. The reaction rate is such that the hydrolysis of acetyl‐CoA is within the linear range. Controls are carried out in absence of enzyme, acetyl‐CoA, or PAS. The amount of CoA produced in the reaction is determined by comparison with a standard curve obtained with DTNB. Third Method. N‐Acetylation of arylamines (PAS) can also be measured by the colorimetric detection of remaining arylamine with 4‐ dimethylaminobenzaldehyde (DMAB) in 96‐well plates as described by Coroneos et al. (1991). This method can be used to measure the activity of recombinant NAT1 and to measure endogenous NAT1 activity in cell extracts. Briefly, NAT1 activity in cell extracts is detected as described previously (Sinclair and Sim, 1997) in a total volume of 100 l. Cell extracts (50 l, obtained from cells exposed or not to oxidants) and PAS (200 M final concentration) in assay buffer (20 mM Tris‐HCl, 1 mM EDTA, pH 7.5)
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are preincubated at 37 for 5 min. AcCoA (400 M final) is added to start the reaction and the samples are incubated at 37 for different times (up to 30 min). The reaction is quenched with 100 l of ice‐cold aqueous TCA (20%, w/v) and proteins are pelleted by centrifugation for 5 min at 12,000g. DMAB [800 l, 5% (w/v) in 9:1 acetonitrile:water] is then added and absorbance is measured in 10‐mm path length cuvettes at 450 nm. The amount of remaining arylamine (not acetylated) is determined from a standard curve. All assays are performed in triplicate, in conditions such that the initial rates are linear. Enzyme activities are normalized according to the protein concentration of cell extracts. Controls are carried out without extract, PAS, or AcCoA. Effect of H2O2 on Recombinant NAT1
As stated earlier, H2O2 is one of the major cellular oxidants. It regulates several cell functions by oxidizing active cysteine residues in proteins (Lee et al., 2002). The physiological/pathophysiological concentration of H2O2 can reach the micromolar range (Halliwell and Gutteridge, 1999), although concentrations above 500 M have been detected (Spector and Garner, 1981). In most studies on the effect of H2O2 on enzyme activities, H2O2 is added as a bolus to the enzyme (either purified or in cell extracts). After an incubation period, residual activity is measured (generally excess H2O2 is removed by treatment with catalase) (Borutaite and Brown, 2001; Caselli et al., 1998; Lee et al., 1998). Using this approach, we showed that recombinant NAT1 is inactivated by H2O2 in a dose‐dependent manner (Atmane et al., 2003). Inactivated enzyme can be reactivated by adding an excess of reducing agents such as DTT or glutathione (GSH) (Atmane et al., 2003). Although simple, the bolus addition of H2O2 to purified enzymes or cell extracts is far from being physiological. Therefore to assess the effect of H2O2 on recombinant NAT1 activity in more realistic and physiological conditions, we used an enzyme system that continuously generates physiological levels of H2O2. The glucose/glucose oxidase or xanthine/ xanthine oxidase systems can be used to this end (Barbouti et al., 2002; Lee et al., 2002; Mueller et al., 1997; Ravid et al., 2002). We used the glucose/ glucose oxidase system (Fig. 3A). We assessed the effect of continuous generation of H2O2 by incubating purified NAT1 (1.5 M) with glucose oxidase (0.15 units/ml, from Aspergillus niger, Sigma) and glucose (5 mM) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA (total volume of 15 l) at 37 . At various time intervals, catalase (300 units/ml, bovine from Sigma) is added and residual NAT1 activity is measured. In controls, catalase (300 unit/ml) is added directly to the glucose/glucose oxidase system. We showed that H2O2 is produced at a constant rate of 6 M H2O2/min in these experiments,
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FIG. 3. (A) Inactivation of human recombinant NAT1 by continuous generation of H2O2 via the glucose/glucose oxidase enzyme system. NAT1 was incubated with glucose (5 mM) and glucose oxidase (0.15 unit/ml) in 25 mM Tris‐HCl, 1 mM EDTA, pH 7.5, at 37 for the indicated times. After the addition of catalase (300 units/ml), residual NAT1 activity was measured. NAT1 was inactivated in a time‐dependent manner. Treatment of NAT1 with the glucose/glucose oxidase system in the presence of catalase did not result in inactivation of the enzyme, showing that NAT1 is indeed inactivated by the generated H2O2. (B) Inactivation of endogenous NAT1 in MCF7 cells upon exposure to peroxynitrite (PN). MCF7 cells in culture dishes were exposed to the specified final concentrations of PN in 10 ml of PBS for 10 min at 37 . After washing with PBS, cells were scraped into lysis buffer, a total extract was made, and NAT1 activity was measured. Endogenous NAT1 was inactivated by PN in a dose‐dependent manner, showing that NAT1 is a target of PN in cells.
which corresponds to physiological levels (Ravid et al., 2002). The amount of H2O2 generated by 0.15 units/ml of glucose oxidase is determined by measuring the absorbance at 240 nm and/or with the peroxidase/orthophenylene‐diamine assay as described previously (Barbouti et al., 2002; Panayiotidis et al., 1999).
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NAT1 is inactivated by the continuous generation of H2O2 in a time‐ dependent manner (Fig. 3A). When exposed to the glucose/glucose oxidase system in the presence of catalase, NAT1 is not inactivated (not shown), demonstrating that the oxidative modification of NAT1 by H2O2 leads to its inactivation. H2O2‐dependent inactivation of NAT1 is reversible as the enzyme is fully active when its oxidized form is incubated with reducing agents (Atmane et al., 2003). The continuous generation of H2O2 by the glucose/glucose oxidase system can also be used to expose cells to H2O2 and to study the subsequent effect of this oxidant on enzymes. We performed such an experiment with cultured lens epithelial cells in petri dishes and showed that endogenous NAT1 and NAT2 (measured in cell extracts) are impaired by continuous exposure to H2O2 (Dairou et al., 2005). A similar protocol using cultured cells and peroxynitrite has been used and is described in more detail here. Effect of Peroxynitrite (PN) on Cellular NAT1
Peroxynitrite is a one of the major biologically relevant oxidants. PN is a highly reactive nitrogen species that exerts many of its biological effects through its capacities to alter protein structure and to function via cysteine oxidation and tyrosine nitration (Groves, 1999). PN is formed during the diffusion rate‐limited bimolecular reaction between nitric oxide and superoxide ion. PN formation and reactions are thought to contribute to the pathogenesis of several physiopathological processes, such as chronic inflammation, sepsis, ischemia‐reperfusion, and cancer (Radi et al., 2001). Physiological concentrations of PN in vivo have been estimated to be around 50 M; however, concentrations of around 500 M have been detected within phagolysosomes of activated macrophages (Denicola et al., 1993). PN has been demonstrated to inhibit enzymatic processes in vitro through either nitration or oxidation of critical amino acids or cofactors (Crow et al., 1995; Mihm and Bauer, 2002; Stachowiak et al., 1998). We assessed the sensitivity of human NAT1 to PN‐dependent inactivation by exposing MCF7 cells (a human breast carcinoma cell line known to express NAT1) to physiological concentrations of PN (Fig. 3B) and assaying residual activity (Dairou et al., 2004). MCF7 cells are cultured as monolayers in 100‐mm petri dishes at 37 in Dulbecco’s modified Eagles medium supplemented with 20% (v/v) fetal bovine serum and penicillin/ streptomycin. At 90% confluence, cell monolayers are washed with phosphate‐buffered saline (PBS) (Ca2þ/Mg2þ). Cell monolayers are exposed to different concentrations of PN (bolus addition of peroxynitrite obtained from Calbiochem‐Novabiochem) in 10 ml of PBS and kept for 10 min at 37 . Controls consist of decomposed PN (obtained by allowing decomposi-
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tion at room temperature in the dark for 48 h) or PBS only. After treatment, monolayers are washed with PBS, scraped in 1 ml of lysis buffer (20 mM Tris‐HCl, 1 mM EDTA, pH 7.5, 0.2 % Triton X‐100, protease inhibitors), and centrifuged for 45 min at 100,000 g. The Bradford method is used to determine the protein concentration of supernatants (total cell extract). Cell extracts are all adjusted to the same protein concentration by adding 20 mM Tris‐HCl, 1 mM EDTA, pH 7.5, and then subjected to the enzymatic assays described earlier. Exposure of cells to PN for 10 min leads to the concentration‐dependent inactivation of endogenous NAT1 (Fig. 3B). Given that PN may be highly toxic for cells, cell viability must be checked using standard protocols (e.g., Trypan blue) to adjust PN concentrations and time of exposure. To mimic physiological PN generation, we have also used 3‐morpholinosydnonimine N‐ethylcarbamide (SIN1), a PN donor. This compound releases both superoxide and nitric oxide at a constant rate, leading to the formation of PN (Groves, 1999; Takakura et al., 1999). SIN1, which is more stable than PN itself, attacks many biological targets in the same manner as PN and has thus been used widely as a source of PN in studies using cultured cells (Singh et al., 1999; Takakura et al., 1999). It is important to remember that the amount of PN generated by SIN1 is lower than the amount of the parent PN donor (Hosker et al., 2001; Percival et al., 1999). Thus, higher concentrations of SIN1 are generally used. Treatment of MCF7 cells with SIN1 gave similar results to those obtained with PN. Decomposed SIN1 may be used to ensure that the observed effect is due to the generation of PN by SIN1. More details about the chemistry of SIN1 can be found in Singh et al. (1999). The inactivation of endogenous NAT1 in cells exposed to PN or SIN1 showed that the reducing intracellular environment of MCF7 cells does not protect endogenous NAT1 sufficiently from PN‐dependent inactivation (Dairou et al., 2004), suggesting that NAT1 is likely to be a target of PN in vivo. Similar results have been obtained with human lens epithelial cells, which are known to have high levels of protective antioxidant systems (Dairou et al., 2005). These results are also supported by data obtained in vitro using recombinant NAT1. Indeed, using the kinetics approach published by Radi et al. (1991), we have shown that NAT1 is inactivated rapidly by PN with a second‐order rate constant (kinact) of 5 104 M1s1. In addition, unlike the H2O2‐dependent inactivation of NAT1, the PN‐ dependent inactivation of NAT1 is not reversed by reducing agents (Dairou et al., 2004). As stated earlier, PN can oxidize cysteine and nitrate tyrosine residues. Chemical modification of recombinant NAT1 by acetylimidazole (a compound that modifies tyrosine side chains) did not inactivate NAT1. Substrate protection assays with AcCoA and CoA were used
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to demonstrate that the oxidative modification of the catalytic cysteine residue of NAT1 by H2O2 or PN was responsible for the inactivation of the enzyme. The active site cysteine residue of NAT is known to form a covalent acetylcysteinyl enzyme with the acetyl group of AcCoA, the physiological acetyl donor substrate of NAT, but not with CoA (the product resulting from hydrolysis of AcCoA) (Pompeo et al., 2002; Riddle and Jencks, 1971). Thus, recombinant NAT1 was preincubated with increasing concentrations of AcCoA and CoA (from 100 M to 5 mM final) and further incubated with a given concentration of PN (or SIN1) that inhibits 90% of NAT1 activity. AcCoA protected NAT1 activity from inactivation by PN in a dose‐dependent manner. Indeed, 73% of the enzyme activity was protected with 5 mM AcCoA, compared to 9% with 5 mM CoA. Although we cannot rule out the possibility that other amino acids of NAT1 are modified by PN, it appears that it is the specific oxidation of the catalytic cysteine residue that leads to enzyme inactivation (Dairou et al., 2004). Overall, these studies show that human NAT1 and, more broadly, NAT enzymes may be the target of biological oxidants. Oxidative‐dependent inactivation of these enzymes could affect the metabolic pathway of numerous xenobiotics. Further studies are needed to assess the real impact of such a regulation in terms of pharmacological susceptibility.
Acknowledgments The authors’ laboratory work was supported by grants from ARC (Association pour la Recherche sur le Cancer), AFM (Association Franc¸ aise contre les Myopathies) and Re´ tina‐ France. J.D. holds a postdoctoral fellowship from AFM. N.A. holds a Ph.D. fellowship from le Ministe`re de la Jeunesse, de l’Education Nationale et de la Recherche.
References Adam, P. J., Berry, J., Loader, J. A., Tyson, K. L., Craggs, G., Smith, P., De Belin, J., Steers, G., Pezzella, F., Sachsenmeir, K. F., Stamps, A. C., Herath, A., Sim, E., O’Hare, M. J., Harris, A. L., and Terrett, J. A. (2003). Arylamine N‐acetyltransferase‐1 is highly expressed in breast cancers and conveys enhanced growth and resistance to etoposide in vitro. Mol. Cancer Res. 1, 826–835. Atmane, N., Dairou, J., Paul, A., Dupret, J. M., and Rodrigues‐Lima, F. (2003). Redox regulation of the human xenobiotic metabolizing enzyme arylamine N‐acetyltransferase 1 (NAT1): Reversible inactivation by hydrogen peroxide. J. Biol. Chem. 278, 35086–35092. Badawi, A. F., Hirvonen, A., Bell, D. A., Lang, N. P., and Kadlubar, F. F. (1995). Role of aromatic amine acetyltransferases, NAT1 and NAT2, in carcinogen‐DNA adduct formation in the human urinary bladder. Cancer Res. 55, 5230–5237.
226
phase ii conjugation enzymes and transport systems
[12]
Barbouti, A., Paschalis‐Thomas, D., Lambros, N., Tenopoulou, M., and Galaris, D. (2002). DNA damage and apoptosis in hydrogen peroxide‐exposed jurkat cells: Bolus addition versus continuous generation of H2O2. Free Radic. Biol. Med. 33, 691–702. Bernassola, F., Rossi, A., and Melino, G. (1999). Regulation of transglutaminase by nitric oxide. Ann. N.Y. Acad. Sci. 887, 83–91. Boelsterli, U. (2003). ‘‘Mechanistic Toxicology: The Molecular Basis of How Chemicals Disrupt Biological Targets.’’ Taylor & Francis, London. Borutaite, V., and Brown, G. (2001). Caspases are reversibly inactivated by hydrogen peroxide. FEBS Lett. 500, 114–118. Brooke, E. W., Davies, S. G., Mulvaney, A. W., Pompeo, F., Sim, E., and Vickers, R. J. (2003). An approach to identifying novel substrates of bacterial arylamine N‐acetyltransferases. Bioorg. Med. Chem. 11, 1227–1234. Butcher, N. J., Arulpragasam, A., and Minchin, R. F. (2004). Proteasomal degradation of N‐acetyltransferase 1 is prevented by acetylation of the active site cysteine: A mechanism for the slow acetylator phenotype and substrate‐dependent down‐regulation. J. Biol. Chem. 279, 22131–22137. Butcher, N. J., Ilett, K. F., and Minchin, R. F. (2000). Substrate‐dependent regulation of human arylamine N‐acetyltransferase‐1 in cultured cells. Mol. Pharmacol. 57, 468–473. Caselli, A., Marzocchini, R., Camici, G., Manao, G., Moneti, G., Pieraccini, G., and Ramponi, G. (1998). The inactivation mechanism of low molecular weight phosphotyrosine‐protein phosphatase by H2O2. J. Biol. Chem. 273, 32554–32560. Coroneos, E., Gordon, J. W., Kelly, S. L., Wang, P. D., and Sim, E. (1991). Drug metabolising N‐acetyltransferase activity in human cell lines. Biochim. Biophys. Acta 1073, 593–599. Crow, J., McCord, J., and Beckman, J. (1995). Sensitivity of the essential zinc‐thiolate moiety of yeast alcohol dehydrogenase to hypochlorite and peroxynitrite. Biochemistry 34, 3544–3552. Dairou, J., Atmane, N., Dupret, J. M., and Rodrigues‐Lima, F. (2003). Reversible inhibition of the human xenobiotic‐metabolizing enzyme arylamine N‐acetyltransferase 1 by S‐nitrosothiols. Biochem. Biophys. Res. Commun. 307, 1059–1065. Dairou, J., Atmane, N., Rodrigues‐Lima, F., and Dupret, J. M. (2004). Peroxynitrite irreversibly inactivates the human xenobiotic‐metabolizing enzyme arylamine N‐acetyltransferase 1 (NAT1) in human breast cancer cells: A cellular and mechanistic study. J. Biol. Chem. 279, 7708–7714. Dairou, J., Malecaze, F., Dupret, J. M., and Rodrigues‐Lima, F. (2005). The xenobiotic‐ metabolizing enzymes arylamine N‐acetyltransferases (NAT) in human lens epithelial cells: Inactivation by cellular oxidants and UVB‐induced oxidative stress. Mol. Pharmacol. 67, 1299–1306. Delome´ nie, C., Fouix, S., Longuemaux, S., Brahimi, N., Bizet, C., Picard, B., Denamur, E., and Dupret, J. M. (2001). Identification and functional characterization of arylamine n‐acetyltransferases in eubacteria: Evidence for highly selective acetylation of 5‐aminosalicylic acid. J. Bacteriol. 183, 3417–3427. Denicola, A., Rubbo, H., Rodriguez, D., and Radi, R. (1993). Peroxynitrite‐mediated cytotoxicity to Trypanosoma cruzi. Arch. Biochem. Biophys. 304, 279–286. Dupret, J. M., Dairou, J., Atmane, N., and Rodrigues‐Lima, F. (2004). Pharmacogenetics, regulation and structural properties of the drug‐metabolizing enzymes arylamine N‐acetyltransferases. Curr. Pharmacogenomics 2, 333–338. Dupret, J. M., and Grant, D. M. (1992). Site‐directed mutagenesis of recombinant human arylamine N‐acetyltransferase expressed in Escherichia coli: Evidence for direct involvement of Cys68 in the catalytic mechanism of polymorphic human NAT2. J. Biol. Chem. 267, 7381–7385.
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Dupret, J. M., and Rodrigues‐Lima, F. (2005). Structure and regulation of the drug‐ metabolizing enzymes arylamine N‐acetyltransferases. Curr. Med. Chem. 12, 763–771. Fretland, A. J., Doll, M. A., Leff, M. A., and Hein, D. W. (2001a). Functional characterization of nucleotide polymorphisms in the coding region of N‐acetyltransferase 1. Pharmacogenetics 11, 511–520. Fretland, A. J., Leff, M. A., Doll, M. A., and Hein, D. W. (2001b). Functional characterization of human N‐acetyltransferase 2 (NAT2) single nucleotide polymorphisms. Pharmacogenetics 11, 207–215. Groves, J. (1999). Peroxynitrite: Reactive, invasive and enigmatic. Curr. Opin. Chem. Biol. 3, 226–235. Guex, N., and Peitsch, M. C. (1997). SWISS‐MODEL and the Swiss‐PdbViewer: An environment for comparative protein modeling. Electrophoresis 18, 2714–2723. Guo, Z., Vath, G. M., Wagner, C. R., and Hanna, P. E. (2003). Arylamine N‐acetyltransferases: Covalent modification and inactivation of hamster NAT1 by bromoacetamido derivatives of aniline and 2‐aminofluorene. J. Protein Chem. 22, 631–642. Guo, Z., Wagner, C. R., and Hanna, P. E. (2004). Mass spectrometric investigation of the mechanism of inactivation of hamster arylamine N‐acetyltransferase 1 by N‐hydroxy‐2‐ acetylaminofluorene. Chem. Res. Toxicol. 17, 275–286. Halliwell, B., and Gutteridge, J. M. C. (1999). ‘‘Free Radicals in Biology and Medicine.’’ Oxford Univ. Press, Oxford. Hein, D. W. (2002). Molecular genetics and function of NAT1 and NAT2: Role in aromatic amine metabolism and carcinogenesis. Mutat. Res. 65, 506–507. Hein, D. W., Doll, M. A., Fretland, A. J., Leff, M. A., Webb, S. J., Xiao, G. H., Devanaboyina, U. S., Nangju, N. A., and Feng, Y. (2000). Molecular genetics and epidemiology of the NAT1 and NAT2 acetylation polymorphisms. Cancer Epidemiol. Biomark. Prev. 9, 29–42. Hickman, D., Pope, J., Patil, S. D., Fakis, G., Smelt, V., Stanley, L. A., Payton, M., Unadkat, J. D., and Sim, E. (1998). Expression of arylamine N‐acetyltransferase in human intestine. Gut 42, 402–409. Holton, S. J., Dairou, J., Sandy, J., Rodrigues‐Lima, F., Dupret, J. M., Noble, M. E. M., and Sim, E. (2005). Structure of Mesorhizobium loti arylamine N‐acetyltransferase 1. Acta Cryst. F61, 14–16. Hosker, B., Bilton, R., Lowe, G., and Loweth, A. (2001). Quantification of peroxynitrite to mixed radical and nitric oxide mediated cell death. Diabetologia 44. [Abstract.] . Husain, A., Barker, D. F., States, J. C., Doll, M. A., and Hein, D. W. (2004). Identification of the major promoter and non‐coding exons of the human arylamine N‐acetyltransferase 1 gene (NAT1). Pharmacogenetics 14, 397–406. Konorev, E. A., Hogg, N., and Kalyanaraman, B. (1998). Rapid and irreversible inhibition of creatine kinase by peroxynitrite. FEBS Lett. 427, 171–174. Lee, S., Kwon, K., Kim, S., and Rhee, S. (1998). Reversible inactivation of protein‐tyrosine phosphatase 1B in A431 cells stimulated with epidermal growth factor. J. Biol. Chem. 273, 15366–15372. Lee, S., Yang, K., Kwon, J., Lee, C., Jeong, W., and Rhee, S. (2002). Reversible inactivation of the tumor suppressor PTEN by H2O2. J. Biol. Chem. 277, 20336–20342. Lin, H., Kent, U., Zhang, H., Waskell, L., and Hollenberg, P. (2003). Mutation of tyrosine 190 to alanine eliminates the inactivation of cytochrome P450 2B1 by peroxynitrite. Chem. Res. Toxicol. 16, 129–136. Liu, L., and Stamler, J. S. (1999). NO: An inhibitor of cell death. Cell Death Differ. 6, 937–942. Matas, N., Thygesen, P., Stacey, M., Risch, A., and Sim, E. (1997). Mapping AAC1, AAC2 and AACP, the genes for arylamine N‐acetyltransferases, carcinogen metabolising
228
phase ii conjugation enzymes and transport systems
[12]
enzymes on human chromosome 8p22, a region frequently deleted in tumours. Cytogenet. Cell Genet. 77, 290–295. Mihm, M., and Bauer, J. (2002). Peroxynitrite‐induced inhibition and nitration of cardiac myofibrillar creatine kinase. Biochimie 84, 1013–1019. Mohr, S., Zech, B., Lapetina, E. G., and Brune, B. (1997). Inhibition of caspase‐3 by S‐nitrosation and oxidation caused by nitric oxide. Biochem. Biophys. Res. Commun. 238, 387–391. Mueller, S., Riedel, H., and Stremmel, W. (1997). Determination of catalase activity at physiological hydrogen peroxide concentrations. Anal. Biochem. 245, 55–60. Pagano, G. (2002). Redox‐modulated xenobiotic action and ROS formation: A mirror or a window? Hum. Exp. Toxicol. 21, 77–81. Panayiotidis, M., Tsolas, O., and Galaris, D. (1999). Glucose oxidase‐produced H2O2 induces Ca2þ‐dependent DNA damage in human peripheral blood lymhocytes. Free Radic. Biol. Med. 26, 548–556. Payton, M., Mushtaq, A., Yu, T. W., Wu, L. J., Sinclair, J., and Sim, E. (2001). Eubacterial arylamine N‐acetyltransferases: Identification and comparison of 18 members of the protein family with conserved active site cysteine, histidine and aspartate residues. Microbiology 147, 1137–1147. Percival, M., Ouellet, M., Campagnolo, C., Claveau, D., and Li, C. (1999). Inhibition of cathepsin K by nitric oxide donors: Evidence for the formation of mixed disulfides and a sulfenic acid. Biochemistry 38, 13574–13583. Pompeo, F., Brooke, E., Akane, K., Mushtaq, A., and Sim, E. (2002). The pharmacogenetics of NAT: Structural aspects. Pharmacogenomics 3, 19–30. Radi, R., Beckman, J., Bush, K., and Freeman, B. (1991). Peroxynitrite oxidation of sulfhydryls. J. Biol. Chem. 266, 4244–4250. Radi, R., Peluffo, G., Alvarez, M. N., Naviliat, M., and Cayota, A. (2001). Unraveling peroxynitrite formation in biological systems. Free Radic. Biol. Med. 30, 463–488. Ravid, T., Sweeney, C., Gee, P., Carraway, K., III, and Goldkorn, T. (2002). Epidermal growth factor receptor activation under oxidative stress fails to promote c‐Cbl mediated down‐regulation. J. Biol. Chem. 277, 31214–31219. Riddle, B., and Jencks, W. P. (1971). Acetyl‐coenzyme A: Arylamine N‐acetyltransferase: Role of the acetyl‐enzyme intermediate and the effects of substituents on the rate. J. Biol. Chem. 246, 3250–3258. Rodrigues‐Lima, F., Cooper, R., Goudeau, B., Atmane, N., Chamagne, A., Butler‐Browne, G., Sim, E., Vicart, P., and Dupret, J. (2003). Skeletal muscles express the xenobiotic‐ metabolizing enzyme arylamine N‐acetyltransferase. J. Histochem. Cytochem. 51, 789–796. Rodrigues‐Lima, F., Delome´ nie, C., Goodfellow, G. H., Grant, D. M., and Dupret, J. M. (2001). Homology modelling and structural analysis of human arylamine N‐acetyltransferase NAT1: Evidence for the conservation of a cysteine protease catalytic domain and an active‐site loop. Biochem. J. 356, 327–334. Rodrigues‐Lima, F., and Dupret, J. M. (2002a). 3D model of human arylamine N‐acetyltransferase 2: Structural basis of the slow acetylator phenotype of the R64Q variant and analysis of the active site loop. Biochem. Biophys. Res. Commun. 291, 116–123. Rodrigues‐Lima, F., and Dupret, J. M. (2002b). In silico sequence analysis of arylamine N‐acetyltransferases: Evidence for an absence of lateral gene transfer from bacteria to vertebrates and first description of paralogs in bacteria. Biochem. Biophys. Res. Commun. 293, 783–792. Rodrigues‐Lima, F., and Dupret, J. M. (2004). Regulation of the activity of the human drug metabolizing enzyme arylamine N‐acetyltransferase 1: Role of genetic and non genetic factors. Curr. Pharm. Des. 10, 2519–2524.
[12]
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Sandy, J., Mushtaq, A., Kawamura, A., Sinclair, J., Noble, M., and Sim, E. (2002). The structure of arylamine N‐acetyltransferase from Mycobacterium smegmatis: An enzyme which inactivates the anti‐tubercular drug, isoniazid. J. Mol. Biol. 318, 1071–1083. Sinclair, J., and Sim, E. (1997). A fragment consisting of the first 204 amino‐terminal amino acids of human arylamine N‐acetyltransferase one (NAT1) and the first transacetylation step of catalysis. Biochem. Pharmacol. 53, 11–16. Sinclair, J. C., Sandy, J., Delgoda, R., Sim, E., and Noble, M. E. (2000). Structure of arylamine N‐acetyltransferase reveals a catalytic triad. Nature Struct. Biol. 7, 560–564. Singh, R. J., Hogg, N., Joseph, J., Konorev, E., and Kalyanaraman, B. (1999). The peroxynitrite generator, SIN‐1, becomes a nitric oxide donor in the presence of electron acceptors. Arch. Biochem. Biophys. 361, 331–339. Spector, A., and Garner, W. H. (1981). Hydrogen peroxide and human cataract. Exp. Eye. Res. 33, 673–681. Stachowiak, O., Dolder, M., Wallimann, T., and Richter, C. (1998). Mitochondrial creatine kinase is a prime target of peroxynitrite‐induced modification and inactivation. J. Biol. Chem. 273, 16694–16699. Takakura, K., Beckman, J., MacMillan‐Crow, L., and Crow, J. (1999). Rapid and irreversible inactivation of protein tyrosine phosphatases PTP1B, CD45, and LAR by peroxynitrite. Arch. Biochem. Biophys. 369, 197–207. Tannickal, V., and Fanburg, B. (2000). Reactive oxygen species in cell signaling. Am. J. Physiol. Lung Cell Mol. Physiol. 279, 1005–1028. Wang, H., Vath, G. M., Gleason, K. J., Hanna, P. E., and Wagner, C. R. (2004). Probing the mechanism of hamster arylamine N‐acetyltransferase 2 acetylation by active site modification, site‐directed mutagenesis and pre‐steady state and steady state kinetic studies. Biochemistry 43, 8234–8246. Westwood, I. M., Holton, S. J., Rodrigues‐Lima, F., Dupret, J.‐M., Bhakta, S., Noble, M. E. M., and Sim, E. (2005). Expression, purification, characterisation and structure of Pseudomonas aeruginosa arylamine N‐acetyltransferase. Biochem. J. 385, 605–612. Wong, P., Eiserich, J., Reddy, S., Lopez, C., Cross, C., and van der Vliet, A. (2001). Inactivation of glutathione S‐transferase by nitric oxide‐derived oxidants: Exploring a role for tyrosine nitration. Arch. Biochem. Biophys. 394, 216–228. Xian, M., Chen, X., Liu, Z., Wang, K., and Wang, P. (2000a). Inhibition of papain by S‐nitrosothiols. J. Biol. Chem. 275, 20467–20473. Xian, M., Wang, K., Chen, X., Hou, Y., McGill, A., Chen, X., Zhou, B., Zhang, Z., Chen, J., and Wang, P. (2000b). Inhibition of protein tyrosine phosphatases by low‐molecular‐ weight S‐nitrosothiols and S‐nitrosylated human serum albumin. Biochem. Biophys. Res. Commun. 268, 310–314.
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Wild, D., Feser, W., Michel, S., Lord, H. L., and Josephy, P. D. (1995). Metabolic activation of heterocyclic aromatic amines catalyzed by human arylamine N‐acetyltransferase isozymes (NAT1 and NAT2) expressed in Salmonella typhimurium. Carcinogenesis 16, 643–648. Xenarios, I., Salwinski, L., Duan, X. J., Higney, P., Kim, S. M., and Eisenberg, D. (2002). DIP, the Database of Interacting Proteins: A research tool for studying cellular networks of protein interactions. Nucleic Acids Res. 30, 303–305. Zhou, H., Josephy, P. D., Kim, D., and Guengerich, F. P. (2004). Functional characterization of four allelic variants of human cytochrome P450 1A2. Arch. Biochem. Biophys. 422, 23–30.
[12] Inactivation of Human Arylamine N‐Acetyltransferase 1 by Hydrogen Peroxide and Peroxynitrite By JEAN‐MARIE DUPRET, JULIEN DAIROU, NOUREDDINE ATMANE , and FERNANDO RODRIGUES‐LIMA Abstract
Arylamine N‐acetyltransferases (NAT) are xenobiotic‐metabolizing enzymes responsible for the acetylation of many arylamine and heterocyclic amines. They therefore play an important role in the detoxification and activation of numerous drugs and carcinogens. Two closely related isoforms (NAT1 and NAT2) have been described in humans. NAT2 is present mainly in the liver and intestine, whereas NAT1 is found in a wide range of tissues. Interindividual variations in NAT genes have been shown to be a potential source of pharmacological and/or pathological susceptibility. Evidence now shows that redox conditions may also contribute to overall NAT activity. This chapter summarizes current knowledge on human NAT1 regulation by reactive oxygen and nitrogen species. Introduction
Arylamine N‐acetyltransferases (NATs) are xenobiotic‐metabolizing enzymes (XME) that catalyze the transfer of an acetyl moiety from acetyl‐CoA to the nitrogen atom of primary arylamines and hydrazines. They are also responsible for the O‐acetylation of N‐hydroxyarylamines (Pompeo et al., 2002). NATs therefore participate in the detoxification and/or activation of a variety of drugs and carcinogens (Badawi et al., 1995; Hein, 2002). NATs have been identified in several species, ranging from bacteria to mammals (Delome´nie et al., 2001; Payton et al., 2001; Rodrigues‐Lima and METHODS IN ENZYMOLOGY, VOL. 400 Copyright 2005, Elsevier Inc. All rights reserved.
0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)00012-1
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Dupret, 2002b). Two functional isoforms of NAT (NAT1 and NAT2) have been described in humans (Matas et al., 1997). NAT1 is found in a wide range of tissues and organs (Rodrigues‐Lima et al., 2003). NAT2 is present mainly in the liver and intestine (Hickman et al., 1998). However, it is likely that NAT2 expression is more widespread (Dairou et al., 2005; Dupret and Rodrigues‐Lima, 2005). Genetically determined interindividual variation in NAT2 content and activity is the basis of the well‐known isoniazid acetylation polymorphism. Polymorphism associated with the NAT1 isoform may also be a source of pharmacological susceptibility (Hein et al., 2000). To date, at least 26 NAT1 alleles and 36 NAT2 alleles have been identified, resulting from numerous single‐nucleotide polymorphisms (Dupret et al., 2004). A detailed description of NAT alleles is available at http://www.louisville.edu/medschool/ pharmacology/NAT.html. Although the relationships between polymorphic substitutions and the activity of variant enzymes have been investigated thoroughly (Fretland et al., 2001a,b), further studies are required to determine the effects of nucleotide variations outside the NAT coding regions and to characterize the enzyme activity of allelic variants more fully. Studies have shown that other mechanisms, such as substrate‐dependent downregulation (Butcher et al., 2000, 2004), the existence of splice variants (Husain et al., 2004), and posttranslational inhibition of NAT activity (Atmane et al., 2003; Dairou et al., 2005), may also contribute to the overall activity of NATs. This chapter summarizes current knowledge on NAT inhibition by reactive oxygen or nitrogen species, consistent with the fundamental role in NAT‐mediated catalysis of a conserved active‐site residue. NAT‐Mediated Catalysis
Results of early steady‐state kinetic studies were consistent with a simple two‐step substituted enzyme (‘‘ping‐pong’’) kinetic mechanism for the NAT reaction (Riddle and Jencks, 1971). Subsequent site‐directed mutagenesis and functional studies have shown that an acetylcysteinyl enzyme is involved in the catalytic process (Dupret and Grant, 1992). Finally, the crystallographic determination of the structure of NATs from prokaryotic species (Salmonella typhimurium, Mycobacterium smegmatis, Pseudomonas aeruginosa, Mesorhizobium loti) (Holton et al., 2005; Sandy et al., 2002; Sinclair et al., 2000; Westwood et al., 2005) and the construction of homology models of human NAT1 and NAT2 (Rodrigues‐Lima and Dupret, 2002a; Rodrigues‐Lima et al., 2001) have revealed structural similarities with cysteine proteases. These studies revealed the existence of a protease‐like catalytic triad (residues Cys 68‐His 107‐Asp 122 in humans) (Fig. 1). The strict conservation of these structural features suggests that
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FIG. 1. Ribbon‐and‐stick representation of the human NAT1 catalytic triad. This figure was created with Deep‐View 3.7b2 (Guex and Peitsch, 1997) using the homology model of the N‐terminal domain of human NAT1 (Rodrigues‐Lima et al., 2001). The hydrogen bonds among cysteine, histidine, and aspartate residues are shown as dotted lines.
NATs have adapted a catalytic mechanism commonly found in cysteine proteases for use in the acetyl‐transfer reaction. Although the exact catalytic mechanism of NAT enzymes is not fully understood, it has been shown to depend on the formation of a thiolate‐ imidazolium ion pair between the triad cysteine and histidine (Wang et al., 2004). The use of chemical modification procedures to probe the reactivity of the active‐site cysteine revealed that it undergoes alkylation upon NAT1 labeling with N‐arylbromoacetamido reagents (Guo et al., 2003). This type of covalent modification can also occur upon NAT1‐catalyzed bioactivation of N‐hydroxy‐2‐acetylaminofluorene and subsequent sulfinamide adduct formation (Guo et al., 2004). In addition to its importance as the primary site of covalent modification in NAT1, Cys 68 has been shown to be the molecular target of reactive oxygen or nitrogen species (Atmane et al., 2003; Dairou et al., 2003, 2004). Oxidative Inhibition of NATs
Several enzymes with a reactive catalytic cysteine residue have been shown to be regulated by the reactive oxygen or nitrogen species (ROS and RNS, respectively) generated during oxidative stress (Halliwell and Gutteridge, 1999). Hydrogen peroxide (H2O2), one of the major cellular
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oxidants, inactivates phosphatases both in vitro and in vivo (Caselli et al., 1998; Lee et al., 1998, 2002). Generally, in vivo and in vitro, H2O2 can oxidize the cysteine residue to give cysteine sulfenic acid (cys‐SOH) or disulfide, which can be reduced back to cysteine by cellular reductants (Halliwell and Gutteridge, 1999). S‐nitrosothiols (RSNO) are reactive nitrogen species known to inactivate cysteine‐containing enzymes, such as papain (Xian et al., 2000a), phosphatases (Xian et al., 2000b), and transglutaminases (Bernassola et al., 1999), either through S‐nitrosylation (formation of an S‐NO) or through formation of a mixed disulfide bond. Both modifications can be reversed by reducing conditions. Peroxynitrite (ONOO) also affects protein function by modifying essential reactive thiols or tyrosine residues (Groves, 1999; Radi et al., 1991). It irreversibly inactivates several enzymes, including creatine kinase (Konorev et al., 1998), caspases (Mohr et al., 1997), and phosphatases (Takakura et al., 1999). XMEs such as cytochrome P450 (Lin et al., 2003) and glutathione S‐transferases (Wong et al., 2001) are also irreversibly inactivated by peroxynitrite. We assessed the ability of reactive oxygen and nitrogen species to inactivate human NAT1. We also investigated the possible reactivation of oxidized NAT1 by reducing agents. Our observations led us to propose that H2O2 and RSNO reversibly inhibit human NAT1, whereas peroxynitrite irreversibly inhibits the enzyme (Rodrigues‐Lima and Dupret, 2004) (Fig. 2).
FIG. 2. Schematic representation of the regulation of human NAT1 by oxidation. The catalytic cysteine residue of the reduced active form of NAT1 is present predominantly as a thiolate anion. This enhances its nucleophilic properties but renders NAT1 susceptible to oxidation. Low levels of H2O2 lead to the reversible oxidation of the catalytic cysteine residue to a sulfenic acid (SOH) with concomitant reversible inactivation of NAT1 activity. When NAT1 is oxidized by S‐nitrosothiols (RSNO), a mixed disulfide (NAT1‐S‐SR) is formed at the catalytic cysteine residue with reversible inactivation of NAT1. Oxidation by low concentrations of peroxynitrite leads to the irreversible inactivation of NAT1 via oxidation of the catalytic cysteine to a sulfinic (SO2H) or sulfonic acid (SO3H).
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Here, we describe in vitro and ex vivo approaches that allow the study of the effect of ROS or RNS on the activity of human NAT1. These reactive species are produced in response to a wide variety of stimuli, such as exposure to chemicals, hormones, and ultraviolet, and following exposure to deleterious physiological/physiopathological conditions, such as apoptosis and inflammation (Groves, 1999; Halliwell and Gutteridge, 1999; Liu and Stamler, 1999; Tannickal and Fanburg, 2000). Excessive production of ROS and RNS is known to participate in the pathological processes of several diseases such as cancer and cataract (Halliwell and Gutteridge, 1999). In addition, these oxidants may affect XME‐dependent biotransformation (Boelsterli, 2003; Pagano, 2002). Thus, the protocols described herein may be used to study the sensitivity of NATs toward oxidative stress induced by various types of stimulus.
General Methods
Expression and Purification of Recombinant Human NAT1 To study the effect of oxidants on recombinant human NAT1, the enzyme is expressed (PET28A vector from Novagen) as a polyhistidine‐ tagged fusion protein (His‐NAT1) in Escherichia coli BL21(DE3) grown for 4 h at 37 in the presence of 0.5 mM isopropyl‐1‐thio‐ ‐D‐galactopyranoside. To purify recombinant NAT1, lysates are loaded onto nickel‐ agarose affinity chromatography columns. Purified His‐NAT1 is reduced by treating with 10 mM dithiothreitol (DTT) (final concentration) for 10 min at 4 prior to dialysis against 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA. SDS–PAGE analysis is carried out at each stage of purification, and protein concentrations are determined using a standard Bradford assay. Cells Expressing NAT1 The effects of oxidants on endogenous cellular NAT1 have been studied using MCF7 (human mammary carcinoma) cells, which are known to express NAT1 (Adam et al., 2003). Other cells have also been used to study the effects of oxidants on endogenous NAT1 (Dairou et al., 2005). Given that the NAT1 gene appears to be expressed ubiquitously, it is likely that most cell types could be used. NAT1 Assays NAT1 activity can be detected easily using spectrophotometric assays. HPLC assays may also be used (Dupret and Grant, 1992). Only
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spectrophotometric assays are described here. These assays can be done easily in the wells of microtiter plates. First Method. NAT1 enzyme activity is determined spectrophotometrically (410 nm) using p‐nitrophenylacetate (PNPA) as the acetyl donor and a NAT1‐specific arylamine substrate such as p‐aminosalicylic acid (PAS) (Musthaq et al., 2002). Briefly, oxidized and unoxidized forms of NAT1 (10–50 l) are assayed in a reaction mixture containing 500 M PAS (final concentration) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA. Reactions are started by adding 125 M PNPA (final concentration). Total reaction volume is 1 ml and final enzyme concentration is usually 10–20 nM. After 10 min at 37 , reactions are quenched by SDS (1% final concentration). p‐Nitrophenol, generated through hydrolysis of PNPA by NAT1 in presence of PAS, is quantified by measuring absorbance at 410 nm with an ELISA plate analyzer. One unit of p‐nitrophenol is defined as the amount of enzyme that gives an absorbance at 410 nm of 0.5 per 10 min/ml. Controls are included without enzyme, PNPA, or PAS. All enzymatic reactions are performed in quadruplicate under conditions where the initial rates are linear. Second Method. The rate of hydrolysis of acetyl‐CoA (first substrate) by NAT1 in the presence of the NAT1‐specific arylamine substrate (PAS) can be determined by detecting the free CoA thiol with 5,50 ‐dithio‐bis 2‐ nitrobenzoic acid (DTNB or Ellman’s reagent) as described previously by Brooke et al. (2003). PAS (500 M final) and samples containing purified NAT1 (0.25 g enzyme) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA buffer are mixed and preincubated (37 , 5 min) in a 96‐well ELISA plate. Acetyl‐ CoA is added to a final concentration of 400 M to start the reaction in a total volume of 100 l. The reaction is quenched with 25 l of 5 mM DTNB in guanidine hydrochloride solution (6.4 M guanidine‐HCl, 0.1 M Tris‐HCl, pH 7.3). The absorbance at 405 nm is measured using an ELISA plate analyzer. The reaction rate is such that the hydrolysis of acetyl‐CoA is within the linear range. Controls are carried out in absence of enzyme, acetyl‐CoA, or PAS. The amount of CoA produced in the reaction is determined by comparison with a standard curve obtained with DTNB. Third Method. N‐Acetylation of arylamines (PAS) can also be measured by the colorimetric detection of remaining arylamine with 4‐ dimethylaminobenzaldehyde (DMAB) in 96‐well plates as described by Coroneos et al. (1991). This method can be used to measure the activity of recombinant NAT1 and to measure endogenous NAT1 activity in cell extracts. Briefly, NAT1 activity in cell extracts is detected as described previously (Sinclair and Sim, 1997) in a total volume of 100 l. Cell extracts (50 l, obtained from cells exposed or not to oxidants) and PAS (200 M final concentration) in assay buffer (20 mM Tris‐HCl, 1 mM EDTA, pH 7.5)
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are preincubated at 37 for 5 min. AcCoA (400 M final) is added to start the reaction and the samples are incubated at 37 for different times (up to 30 min). The reaction is quenched with 100 l of ice‐cold aqueous TCA (20%, w/v) and proteins are pelleted by centrifugation for 5 min at 12,000g. DMAB [800 l, 5% (w/v) in 9:1 acetonitrile:water] is then added and absorbance is measured in 10‐mm path length cuvettes at 450 nm. The amount of remaining arylamine (not acetylated) is determined from a standard curve. All assays are performed in triplicate, in conditions such that the initial rates are linear. Enzyme activities are normalized according to the protein concentration of cell extracts. Controls are carried out without extract, PAS, or AcCoA. Effect of H2O2 on Recombinant NAT1
As stated earlier, H2O2 is one of the major cellular oxidants. It regulates several cell functions by oxidizing active cysteine residues in proteins (Lee et al., 2002). The physiological/pathophysiological concentration of H2O2 can reach the micromolar range (Halliwell and Gutteridge, 1999), although concentrations above 500 M have been detected (Spector and Garner, 1981). In most studies on the effect of H2O2 on enzyme activities, H2O2 is added as a bolus to the enzyme (either purified or in cell extracts). After an incubation period, residual activity is measured (generally excess H2O2 is removed by treatment with catalase) (Borutaite and Brown, 2001; Caselli et al., 1998; Lee et al., 1998). Using this approach, we showed that recombinant NAT1 is inactivated by H2O2 in a dose‐dependent manner (Atmane et al., 2003). Inactivated enzyme can be reactivated by adding an excess of reducing agents such as DTT or glutathione (GSH) (Atmane et al., 2003). Although simple, the bolus addition of H2O2 to purified enzymes or cell extracts is far from being physiological. Therefore to assess the effect of H2O2 on recombinant NAT1 activity in more realistic and physiological conditions, we used an enzyme system that continuously generates physiological levels of H2O2. The glucose/glucose oxidase or xanthine/ xanthine oxidase systems can be used to this end (Barbouti et al., 2002; Lee et al., 2002; Mueller et al., 1997; Ravid et al., 2002). We used the glucose/ glucose oxidase system (Fig. 3A). We assessed the effect of continuous generation of H2O2 by incubating purified NAT1 (1.5 M) with glucose oxidase (0.15 units/ml, from Aspergillus niger, Sigma) and glucose (5 mM) in 25 mM Tris‐HCl, pH 7.5, 1 mM EDTA (total volume of 15 l) at 37 . At various time intervals, catalase (300 units/ml, bovine from Sigma) is added and residual NAT1 activity is measured. In controls, catalase (300 unit/ml) is added directly to the glucose/glucose oxidase system. We showed that H2O2 is produced at a constant rate of 6 M H2O2/min in these experiments,
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FIG. 3. (A) Inactivation of human recombinant NAT1 by continuous generation of H2O2 via the glucose/glucose oxidase enzyme system. NAT1 was incubated with glucose (5 mM) and glucose oxidase (0.15 unit/ml) in 25 mM Tris‐HCl, 1 mM EDTA, pH 7.5, at 37 for the indicated times. After the addition of catalase (300 units/ml), residual NAT1 activity was measured. NAT1 was inactivated in a time‐dependent manner. Treatment of NAT1 with the glucose/glucose oxidase system in the presence of catalase did not result in inactivation of the enzyme, showing that NAT1 is indeed inactivated by the generated H2O2. (B) Inactivation of endogenous NAT1 in MCF7 cells upon exposure to peroxynitrite (PN). MCF7 cells in culture dishes were exposed to the specified final concentrations of PN in 10 ml of PBS for 10 min at 37 . After washing with PBS, cells were scraped into lysis buffer, a total extract was made, and NAT1 activity was measured. Endogenous NAT1 was inactivated by PN in a dose‐dependent manner, showing that NAT1 is a target of PN in cells.
which corresponds to physiological levels (Ravid et al., 2002). The amount of H2O2 generated by 0.15 units/ml of glucose oxidase is determined by measuring the absorbance at 240 nm and/or with the peroxidase/orthophenylene‐diamine assay as described previously (Barbouti et al., 2002; Panayiotidis et al., 1999).
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NAT1 is inactivated by the continuous generation of H2O2 in a time‐ dependent manner (Fig. 3A). When exposed to the glucose/glucose oxidase system in the presence of catalase, NAT1 is not inactivated (not shown), demonstrating that the oxidative modification of NAT1 by H2O2 leads to its inactivation. H2O2‐dependent inactivation of NAT1 is reversible as the enzyme is fully active when its oxidized form is incubated with reducing agents (Atmane et al., 2003). The continuous generation of H2O2 by the glucose/glucose oxidase system can also be used to expose cells to H2O2 and to study the subsequent effect of this oxidant on enzymes. We performed such an experiment with cultured lens epithelial cells in petri dishes and showed that endogenous NAT1 and NAT2 (measured in cell extracts) are impaired by continuous exposure to H2O2 (Dairou et al., 2005). A similar protocol using cultured cells and peroxynitrite has been used and is described in more detail here. Effect of Peroxynitrite (PN) on Cellular NAT1
Peroxynitrite is a one of the major biologically relevant oxidants. PN is a highly reactive nitrogen species that exerts many of its biological effects through its capacities to alter protein structure and to function via cysteine oxidation and tyrosine nitration (Groves, 1999). PN is formed during the diffusion rate‐limited bimolecular reaction between nitric oxide and superoxide ion. PN formation and reactions are thought to contribute to the pathogenesis of several physiopathological processes, such as chronic inflammation, sepsis, ischemia‐reperfusion, and cancer (Radi et al., 2001). Physiological concentrations of PN in vivo have been estimated to be around 50 M; however, concentrations of around 500 M have been detected within phagolysosomes of activated macrophages (Denicola et al., 1993). PN has been demonstrated to inhibit enzymatic processes in vitro through either nitration or oxidation of critical amino acids or cofactors (Crow et al., 1995; Mihm and Bauer, 2002; Stachowiak et al., 1998). We assessed the sensitivity of human NAT1 to PN‐dependent inactivation by exposing MCF7 cells (a human breast carcinoma cell line known to express NAT1) to physiological concentrations of PN (Fig. 3B) and assaying residual activity (Dairou et al., 2004). MCF7 cells are cultured as monolayers in 100‐mm petri dishes at 37 in Dulbecco’s modified Eagles medium supplemented with 20% (v/v) fetal bovine serum and penicillin/ streptomycin. At 90% confluence, cell monolayers are washed with phosphate‐buffered saline (PBS) (Ca2þ/Mg2þ). Cell monolayers are exposed to different concentrations of PN (bolus addition of peroxynitrite obtained from Calbiochem‐Novabiochem) in 10 ml of PBS and kept for 10 min at 37 . Controls consist of decomposed PN (obtained by allowing decomposi-
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tion at room temperature in the dark for 48 h) or PBS only. After treatment, monolayers are washed with PBS, scraped in 1 ml of lysis buffer (20 mM Tris‐HCl, 1 mM EDTA, pH 7.5, 0.2 % Triton X‐100, protease inhibitors), and centrifuged for 45 min at 100,000 g. The Bradford method is used to determine the protein concentration of supernatants (total cell extract). Cell extracts are all adjusted to the same protein concentration by adding 20 mM Tris‐HCl, 1 mM EDTA, pH 7.5, and then subjected to the enzymatic assays described earlier. Exposure of cells to PN for 10 min leads to the concentration‐dependent inactivation of endogenous NAT1 (Fig. 3B). Given that PN may be highly toxic for cells, cell viability must be checked using standard protocols (e.g., Trypan blue) to adjust PN concentrations and time of exposure. To mimic physiological PN generation, we have also used 3‐morpholinosydnonimine N‐ethylcarbamide (SIN1), a PN donor. This compound releases both superoxide and nitric oxide at a constant rate, leading to the formation of PN (Groves, 1999; Takakura et al., 1999). SIN1, which is more stable than PN itself, attacks many biological targets in the same manner as PN and has thus been used widely as a source of PN in studies using cultured cells (Singh et al., 1999; Takakura et al., 1999). It is important to remember that the amount of PN generated by SIN1 is lower than the amount of the parent PN donor (Hosker et al., 2001; Percival et al., 1999). Thus, higher concentrations of SIN1 are generally used. Treatment of MCF7 cells with SIN1 gave similar results to those obtained with PN. Decomposed SIN1 may be used to ensure that the observed effect is due to the generation of PN by SIN1. More details about the chemistry of SIN1 can be found in Singh et al. (1999). The inactivation of endogenous NAT1 in cells exposed to PN or SIN1 showed that the reducing intracellular environment of MCF7 cells does not protect endogenous NAT1 sufficiently from PN‐dependent inactivation (Dairou et al., 2004), suggesting that NAT1 is likely to be a target of PN in vivo. Similar results have been obtained with human lens epithelial cells, which are known to have high levels of protective antioxidant systems (Dairou et al., 2005). These results are also supported by data obtained in vitro using recombinant NAT1. Indeed, using the kinetics approach published by Radi et al. (1991), we have shown that NAT1 is inactivated rapidly by PN with a second‐order rate constant (kinact) of 5 104 M1s1. In addition, unlike the H2O2‐dependent inactivation of NAT1, the PN‐ dependent inactivation of NAT1 is not reversed by reducing agents (Dairou et al., 2004). As stated earlier, PN can oxidize cysteine and nitrate tyrosine residues. Chemical modification of recombinant NAT1 by acetylimidazole (a compound that modifies tyrosine side chains) did not inactivate NAT1. Substrate protection assays with AcCoA and CoA were used
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to demonstrate that the oxidative modification of the catalytic cysteine residue of NAT1 by H2O2 or PN was responsible for the inactivation of the enzyme. The active site cysteine residue of NAT is known to form a covalent acetylcysteinyl enzyme with the acetyl group of AcCoA, the physiological acetyl donor substrate of NAT, but not with CoA (the product resulting from hydrolysis of AcCoA) (Pompeo et al., 2002; Riddle and Jencks, 1971). Thus, recombinant NAT1 was preincubated with increasing concentrations of AcCoA and CoA (from 100 M to 5 mM final) and further incubated with a given concentration of PN (or SIN1) that inhibits 90% of NAT1 activity. AcCoA protected NAT1 activity from inactivation by PN in a dose‐dependent manner. Indeed, 73% of the enzyme activity was protected with 5 mM AcCoA, compared to 9% with 5 mM CoA. Although we cannot rule out the possibility that other amino acids of NAT1 are modified by PN, it appears that it is the specific oxidation of the catalytic cysteine residue that leads to enzyme inactivation (Dairou et al., 2004). Overall, these studies show that human NAT1 and, more broadly, NAT enzymes may be the target of biological oxidants. Oxidative‐dependent inactivation of these enzymes could affect the metabolic pathway of numerous xenobiotics. Further studies are needed to assess the real impact of such a regulation in terms of pharmacological susceptibility.
Acknowledgments The authors’ laboratory work was supported by grants from ARC (Association pour la Recherche sur le Cancer), AFM (Association Franc¸ aise contre les Myopathies) and Re´ tina‐ France. J.D. holds a postdoctoral fellowship from AFM. N.A. holds a Ph.D. fellowship from le Ministe`re de la Jeunesse, de l’Education Nationale et de la Recherche.
References Adam, P. J., Berry, J., Loader, J. A., Tyson, K. L., Craggs, G., Smith, P., De Belin, J., Steers, G., Pezzella, F., Sachsenmeir, K. F., Stamps, A. C., Herath, A., Sim, E., O’Hare, M. J., Harris, A. L., and Terrett, J. A. (2003). Arylamine N‐acetyltransferase‐1 is highly expressed in breast cancers and conveys enhanced growth and resistance to etoposide in vitro. Mol. Cancer Res. 1, 826–835. Atmane, N., Dairou, J., Paul, A., Dupret, J. M., and Rodrigues‐Lima, F. (2003). Redox regulation of the human xenobiotic metabolizing enzyme arylamine N‐acetyltransferase 1 (NAT1): Reversible inactivation by hydrogen peroxide. J. Biol. Chem. 278, 35086–35092. Badawi, A. F., Hirvonen, A., Bell, D. A., Lang, N. P., and Kadlubar, F. F. (1995). Role of aromatic amine acetyltransferases, NAT1 and NAT2, in carcinogen‐DNA adduct formation in the human urinary bladder. Cancer Res. 55, 5230–5237.
226
phase ii conjugation enzymes and transport systems
[12]
Barbouti, A., Paschalis‐Thomas, D., Lambros, N., Tenopoulou, M., and Galaris, D. (2002). DNA damage and apoptosis in hydrogen peroxide‐exposed jurkat cells: Bolus addition versus continuous generation of H2O2. Free Radic. Biol. Med. 33, 691–702. Bernassola, F., Rossi, A., and Melino, G. (1999). Regulation of transglutaminase by nitric oxide. Ann. N.Y. Acad. Sci. 887, 83–91. Boelsterli, U. (2003). ‘‘Mechanistic Toxicology: The Molecular Basis of How Chemicals Disrupt Biological Targets.’’ Taylor & Francis, London. Borutaite, V., and Brown, G. (2001). Caspases are reversibly inactivated by hydrogen peroxide. FEBS Lett. 500, 114–118. Brooke, E. W., Davies, S. G., Mulvaney, A. W., Pompeo, F., Sim, E., and Vickers, R. J. (2003). An approach to identifying novel substrates of bacterial arylamine N‐acetyltransferases. Bioorg. Med. Chem. 11, 1227–1234. Butcher, N. J., Arulpragasam, A., and Minchin, R. F. (2004). Proteasomal degradation of N‐acetyltransferase 1 is prevented by acetylation of the active site cysteine: A mechanism for the slow acetylator phenotype and substrate‐dependent down‐regulation. J. Biol. Chem. 279, 22131–22137. Butcher, N. J., Ilett, K. F., and Minchin, R. F. (2000). Substrate‐dependent regulation of human arylamine N‐acetyltransferase‐1 in cultured cells. Mol. Pharmacol. 57, 468–473. Caselli, A., Marzocchini, R., Camici, G., Manao, G., Moneti, G., Pieraccini, G., and Ramponi, G. (1998). The inactivation mechanism of low molecular weight phosphotyrosine‐protein phosphatase by H2O2. J. Biol. Chem. 273, 32554–32560. Coroneos, E., Gordon, J. W., Kelly, S. L., Wang, P. D., and Sim, E. (1991). Drug metabolising N‐acetyltransferase activity in human cell lines. Biochim. Biophys. Acta 1073, 593–599. Crow, J., McCord, J., and Beckman, J. (1995). Sensitivity of the essential zinc‐thiolate moiety of yeast alcohol dehydrogenase to hypochlorite and peroxynitrite. Biochemistry 34, 3544–3552. Dairou, J., Atmane, N., Dupret, J. M., and Rodrigues‐Lima, F. (2003). Reversible inhibition of the human xenobiotic‐metabolizing enzyme arylamine N‐acetyltransferase 1 by S‐nitrosothiols. Biochem. Biophys. Res. Commun. 307, 1059–1065. Dairou, J., Atmane, N., Rodrigues‐Lima, F., and Dupret, J. M. (2004). Peroxynitrite irreversibly inactivates the human xenobiotic‐metabolizing enzyme arylamine N‐acetyltransferase 1 (NAT1) in human breast cancer cells: A cellular and mechanistic study. J. Biol. Chem. 279, 7708–7714. Dairou, J., Malecaze, F., Dupret, J. M., and Rodrigues‐Lima, F. (2005). The xenobiotic‐ metabolizing enzymes arylamine N‐acetyltransferases (NAT) in human lens epithelial cells: Inactivation by cellular oxidants and UVB‐induced oxidative stress. Mol. Pharmacol. 67, 1299–1306. Delome´ nie, C., Fouix, S., Longuemaux, S., Brahimi, N., Bizet, C., Picard, B., Denamur, E., and Dupret, J. M. (2001). Identification and functional characterization of arylamine n‐acetyltransferases in eubacteria: Evidence for highly selective acetylation of 5‐aminosalicylic acid. J. Bacteriol. 183, 3417–3427. Denicola, A., Rubbo, H., Rodriguez, D., and Radi, R. (1993). Peroxynitrite‐mediated cytotoxicity to Trypanosoma cruzi. Arch. Biochem. Biophys. 304, 279–286. Dupret, J. M., Dairou, J., Atmane, N., and Rodrigues‐Lima, F. (2004). Pharmacogenetics, regulation and structural properties of the drug‐metabolizing enzymes arylamine N‐acetyltransferases. Curr. Pharmacogenomics 2, 333–338. Dupret, J. M., and Grant, D. M. (1992). Site‐directed mutagenesis of recombinant human arylamine N‐acetyltransferase expressed in Escherichia coli: Evidence for direct involvement of Cys68 in the catalytic mechanism of polymorphic human NAT2. J. Biol. Chem. 267, 7381–7385.
[12]
oxidative inactivation of NAT1 enzyme
227
Dupret, J. M., and Rodrigues‐Lima, F. (2005). Structure and regulation of the drug‐ metabolizing enzymes arylamine N‐acetyltransferases. Curr. Med. Chem. 12, 763–771. Fretland, A. J., Doll, M. A., Leff, M. A., and Hein, D. W. (2001a). Functional characterization of nucleotide polymorphisms in the coding region of N‐acetyltransferase 1. Pharmacogenetics 11, 511–520. Fretland, A. J., Leff, M. A., Doll, M. A., and Hein, D. W. (2001b). Functional characterization of human N‐acetyltransferase 2 (NAT2) single nucleotide polymorphisms. Pharmacogenetics 11, 207–215. Groves, J. (1999). Peroxynitrite: Reactive, invasive and enigmatic. Curr. Opin. Chem. Biol. 3, 226–235. Guex, N., and Peitsch, M. C. (1997). SWISS‐MODEL and the Swiss‐PdbViewer: An environment for comparative protein modeling. Electrophoresis 18, 2714–2723. Guo, Z., Vath, G. M., Wagner, C. R., and Hanna, P. E. (2003). Arylamine N‐acetyltransferases: Covalent modification and inactivation of hamster NAT1 by bromoacetamido derivatives of aniline and 2‐aminofluorene. J. Protein Chem. 22, 631–642. Guo, Z., Wagner, C. R., and Hanna, P. E. (2004). Mass spectrometric investigation of the mechanism of inactivation of hamster arylamine N‐acetyltransferase 1 by N‐hydroxy‐2‐ acetylaminofluorene. Chem. Res. Toxicol. 17, 275–286. Halliwell, B., and Gutteridge, J. M. C. (1999). ‘‘Free Radicals in Biology and Medicine.’’ Oxford Univ. Press, Oxford. Hein, D. W. (2002). Molecular genetics and function of NAT1 and NAT2: Role in aromatic amine metabolism and carcinogenesis. Mutat. Res. 65, 506–507. Hein, D. W., Doll, M. A., Fretland, A. J., Leff, M. A., Webb, S. J., Xiao, G. H., Devanaboyina, U. S., Nangju, N. A., and Feng, Y. (2000). Molecular genetics and epidemiology of the NAT1 and NAT2 acetylation polymorphisms. Cancer Epidemiol. Biomark. Prev. 9, 29–42. Hickman, D., Pope, J., Patil, S. D., Fakis, G., Smelt, V., Stanley, L. A., Payton, M., Unadkat, J. D., and Sim, E. (1998). Expression of arylamine N‐acetyltransferase in human intestine. Gut 42, 402–409. Holton, S. J., Dairou, J., Sandy, J., Rodrigues‐Lima, F., Dupret, J. M., Noble, M. E. M., and Sim, E. (2005). Structure of Mesorhizobium loti arylamine N‐acetyltransferase 1. Acta Cryst. F61, 14–16. Hosker, B., Bilton, R., Lowe, G., and Loweth, A. (2001). Quantification of peroxynitrite to mixed radical and nitric oxide mediated cell death. Diabetologia 44. [Abstract.] . Husain, A., Barker, D. F., States, J. C., Doll, M. A., and Hein, D. W. (2004). Identification of the major promoter and non‐coding exons of the human arylamine N‐acetyltransferase 1 gene (NAT1). Pharmacogenetics 14, 397–406. Konorev, E. A., Hogg, N., and Kalyanaraman, B. (1998). Rapid and irreversible inhibition of creatine kinase by peroxynitrite. FEBS Lett. 427, 171–174. Lee, S., Kwon, K., Kim, S., and Rhee, S. (1998). Reversible inactivation of protein‐tyrosine phosphatase 1B in A431 cells stimulated with epidermal growth factor. J. Biol. Chem. 273, 15366–15372. Lee, S., Yang, K., Kwon, J., Lee, C., Jeong, W., and Rhee, S. (2002). Reversible inactivation of the tumor suppressor PTEN by H2O2. J. Biol. Chem. 277, 20336–20342. Lin, H., Kent, U., Zhang, H., Waskell, L., and Hollenberg, P. (2003). Mutation of tyrosine 190 to alanine eliminates the inactivation of cytochrome P450 2B1 by peroxynitrite. Chem. Res. Toxicol. 16, 129–136. Liu, L., and Stamler, J. S. (1999). NO: An inhibitor of cell death. Cell Death Differ. 6, 937–942. Matas, N., Thygesen, P., Stacey, M., Risch, A., and Sim, E. (1997). Mapping AAC1, AAC2 and AACP, the genes for arylamine N‐acetyltransferases, carcinogen metabolising
228
phase ii conjugation enzymes and transport systems
[12]
enzymes on human chromosome 8p22, a region frequently deleted in tumours. Cytogenet. Cell Genet. 77, 290–295. Mihm, M., and Bauer, J. (2002). Peroxynitrite‐induced inhibition and nitration of cardiac myofibrillar creatine kinase. Biochimie 84, 1013–1019. Mohr, S., Zech, B., Lapetina, E. G., and Brune, B. (1997). Inhibition of caspase‐3 by S‐nitrosation and oxidation caused by nitric oxide. Biochem. Biophys. Res. Commun. 238, 387–391. Mueller, S., Riedel, H., and Stremmel, W. (1997). Determination of catalase activity at physiological hydrogen peroxide concentrations. Anal. Biochem. 245, 55–60. Pagano, G. (2002). Redox‐modulated xenobiotic action and ROS formation: A mirror or a window? Hum. Exp. Toxicol. 21, 77–81. Panayiotidis, M., Tsolas, O., and Galaris, D. (1999). Glucose oxidase‐produced H2O2 induces Ca2þ‐dependent DNA damage in human peripheral blood lymhocytes. Free Radic. Biol. Med. 26, 548–556. Payton, M., Mushtaq, A., Yu, T. W., Wu, L. J., Sinclair, J., and Sim, E. (2001). Eubacterial arylamine N‐acetyltransferases: Identification and comparison of 18 members of the protein family with conserved active site cysteine, histidine and aspartate residues. Microbiology 147, 1137–1147. Percival, M., Ouellet, M., Campagnolo, C., Claveau, D., and Li, C. (1999). Inhibition of cathepsin K by nitric oxide donors: Evidence for the formation of mixed disulfides and a sulfenic acid. Biochemistry 38, 13574–13583. Pompeo, F., Brooke, E., Akane, K., Mushtaq, A., and Sim, E. (2002). The pharmacogenetics of NAT: Structural aspects. Pharmacogenomics 3, 19–30. Radi, R., Beckman, J., Bush, K., and Freeman, B. (1991). Peroxynitrite oxidation of sulfhydryls. J. Biol. Chem. 266, 4244–4250. Radi, R., Peluffo, G., Alvarez, M. N., Naviliat, M., and Cayota, A. (2001). Unraveling peroxynitrite formation in biological systems. Free Radic. Biol. Med. 30, 463–488. Ravid, T., Sweeney, C., Gee, P., Carraway, K., III, and Goldkorn, T. (2002). Epidermal growth factor receptor activation under oxidative stress fails to promote c‐Cbl mediated down‐regulation. J. Biol. Chem. 277, 31214–31219. Riddle, B., and Jencks, W. P. (1971). Acetyl‐coenzyme A: Arylamine N‐acetyltransferase: Role of the acetyl‐enzyme intermediate and the effects of substituents on the rate. J. Biol. Chem. 246, 3250–3258. Rodrigues‐Lima, F., Cooper, R., Goudeau, B., Atmane, N., Chamagne, A., Butler‐Browne, G., Sim, E., Vicart, P., and Dupret, J. (2003). Skeletal muscles express the xenobiotic‐ metabolizing enzyme arylamine N‐acetyltransferase. J. Histochem. Cytochem. 51, 789–796. Rodrigues‐Lima, F., Delome´ nie, C., Goodfellow, G. H., Grant, D. M., and Dupret, J. M. (2001). Homology modelling and structural analysis of human arylamine N‐acetyltransferase NAT1: Evidence for the conservation of a cysteine protease catalytic domain and an active‐site loop. Biochem. J. 356, 327–334. Rodrigues‐Lima, F., and Dupret, J. M. (2002a). 3D model of human arylamine N‐acetyltransferase 2: Structural basis of the slow acetylator phenotype of the R64Q variant and analysis of the active site loop. Biochem. Biophys. Res. Commun. 291, 116–123. Rodrigues‐Lima, F., and Dupret, J. M. (2002b). In silico sequence analysis of arylamine N‐acetyltransferases: Evidence for an absence of lateral gene transfer from bacteria to vertebrates and first description of paralogs in bacteria. Biochem. Biophys. Res. Commun. 293, 783–792. Rodrigues‐Lima, F., and Dupret, J. M. (2004). Regulation of the activity of the human drug metabolizing enzyme arylamine N‐acetyltransferase 1: Role of genetic and non genetic factors. Curr. Pharm. Des. 10, 2519–2524.
[12]
oxidative inactivation of NAT1 enzyme
229
Sandy, J., Mushtaq, A., Kawamura, A., Sinclair, J., Noble, M., and Sim, E. (2002). The structure of arylamine N‐acetyltransferase from Mycobacterium smegmatis: An enzyme which inactivates the anti‐tubercular drug, isoniazid. J. Mol. Biol. 318, 1071–1083. Sinclair, J., and Sim, E. (1997). A fragment consisting of the first 204 amino‐terminal amino acids of human arylamine N‐acetyltransferase one (NAT1) and the first transacetylation step of catalysis. Biochem. Pharmacol. 53, 11–16. Sinclair, J. C., Sandy, J., Delgoda, R., Sim, E., and Noble, M. E. (2000). Structure of arylamine N‐acetyltransferase reveals a catalytic triad. Nature Struct. Biol. 7, 560–564. Singh, R. J., Hogg, N., Joseph, J., Konorev, E., and Kalyanaraman, B. (1999). The peroxynitrite generator, SIN‐1, becomes a nitric oxide donor in the presence of electron acceptors. Arch. Biochem. Biophys. 361, 331–339. Spector, A., and Garner, W. H. (1981). Hydrogen peroxide and human cataract. Exp. Eye. Res. 33, 673–681. Stachowiak, O., Dolder, M., Wallimann, T., and Richter, C. (1998). Mitochondrial creatine kinase is a prime target of peroxynitrite‐induced modification and inactivation. J. Biol. Chem. 273, 16694–16699. Takakura, K., Beckman, J., MacMillan‐Crow, L., and Crow, J. (1999). Rapid and irreversible inactivation of protein tyrosine phosphatases PTP1B, CD45, and LAR by peroxynitrite. Arch. Biochem. Biophys. 369, 197–207. Tannickal, V., and Fanburg, B. (2000). Reactive oxygen species in cell signaling. Am. J. Physiol. Lung Cell Mol. Physiol. 279, 1005–1028. Wang, H., Vath, G. M., Gleason, K. J., Hanna, P. E., and Wagner, C. R. (2004). Probing the mechanism of hamster arylamine N‐acetyltransferase 2 acetylation by active site modification, site‐directed mutagenesis and pre‐steady state and steady state kinetic studies. Biochemistry 43, 8234–8246. Westwood, I. M., Holton, S. J., Rodrigues‐Lima, F., Dupret, J.‐M., Bhakta, S., Noble, M. E. M., and Sim, E. (2005). Expression, purification, characterisation and structure of Pseudomonas aeruginosa arylamine N‐acetyltransferase. Biochem. J. 385, 605–612. Wong, P., Eiserich, J., Reddy, S., Lopez, C., Cross, C., and van der Vliet, A. (2001). Inactivation of glutathione S‐transferase by nitric oxide‐derived oxidants: Exploring a role for tyrosine nitration. Arch. Biochem. Biophys. 394, 216–228. Xian, M., Chen, X., Liu, Z., Wang, K., and Wang, P. (2000a). Inhibition of papain by S‐nitrosothiols. J. Biol. Chem. 275, 20467–20473. Xian, M., Wang, K., Chen, X., Hou, Y., McGill, A., Chen, X., Zhou, B., Zhang, Z., Chen, J., and Wang, P. (2000b). Inhibition of protein tyrosine phosphatases by low‐molecular‐ weight S‐nitrosothiols and S‐nitrosylated human serum albumin. Biochem. Biophys. Res. Commun. 268, 310–314.