Biochemical Pharmacology xxx (2017) xxx–xxx
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Identification of enzymes responsible for nitrazepam metabolism and toxicity in human Keigo Konishi, Tatsuki Fukami ⇑, Saki Gotoh, Miki Nakajima Drug Metabolism and Toxicology, Faculty of Pharmaceutical Sciences, Kanazawa University, Kanazawa, Japan
a r t i c l e
i n f o
Article history: Received 14 April 2017 Accepted 7 June 2017 Available online xxxx Keywords: AOX1 NAT2 AADAC CYP3A4 Reactive metabolite
a b s t r a c t Nitrazepam (NZP) is a hypnotic agent that rarely causes liver injuries in humans and teratogenicity in rodents. In humans, NZP is primarily metabolized to 7-aminonitrazepam (ANZP) by reduction and subsequently to 7-acetylamino nitrazepam (AANZP) by acetylation. ANZP can be regenerated from AANZP by hydrolysis in rodents, but it is still unclear whether this reaction occurs in humans. In rodents, AANZP may be associated with teratogenicity, while in humans, it is known that drug-induced liver injuries may be caused by NZP reactive metabolite(s). In this study, we attempted to identify the enzymes responsible for NZP metabolism to obtain a basic understanding of this process and the associated metabolite toxicities. We found that the NZP reductase activity in human liver cytosol (HLC) was higher than that in human liver microsomes (HLM). We purified the responsible enzyme(s) from HLC and found that the NZP reductase was aldehyde oxidase 1 (AOX1). The role of AOX1 was confirmed by an observed increase in the NZP reductase activity upon addition of N1-methylnicotinamide, an electron donor of AOX1, as well as inhibition of this activity in HLC in the presence of AOX1 inhibitors. ANZP was acetylated to form AANZP by N-acetyltransferase (NAT) 2. An experiment using recombinant esterases in an inhibition study using HLM revealed that AANZP is hydrolyzed by arylacetamide deacetylase (AADAC) in the human liver. N-Hydroxylamino NZP, which is suspected to be a reactive metabolite, was detected as a conjugate with N-acetyl-L-cysteine through NZP reduction and ANZP hydroxylation reactions. In the latter reaction, the conjugate was readily formed by recombinant CYP3A4 among the various P450 isoforms tested. In sum, we found that AOX1, NAT2, AADAC, and CYP3A4 are the determinants for the pharmacokinetics of NZP and that they confer interindividual variability in sensitivity to NZP side effects. Ó 2017 Published by Elsevier Inc.
1. Introduction Nitrazepam (NZP) is a long-acting benzodiazepine hypnotic agent that is used as an effective treatment for insomnia [1]. Insomnia is referred to as a disorder of initiating or maintaining sleep, and its prevalence in older people is estimated to be 23– 34% [2,3]. NZP works as a hypnotic to prolong sleeping time and decrease the time needed to fall asleep [3], but several side effects, such as liver injury and teratogenicity, can occur. Cholestatic liver failure has been reported as a result of NZP usage [4,5]. These
adverse reactions are thought to be induced through the metabolism of NZP [6,7]. In humans, NZP is primarily metabolized to 7-amino nitrazepam (ANZP) by nitro-reductase(s) and subsequently to 7-acetylamino nitrazepam (AANZP) by N-acetyltransferase(s) (Fig. 1) [8]. In rodents, AANZP is hydrolyzed to ANZP in the liver, but it has not been determined whether this reaction occurs in humans [7]. Within 24 hours of an oral administration of 10 mg NZP in humans, urinary excretion of ANZP and AANZP represented 4.1% and 12.5% of the initial dose, respectively, [9]. Other possible NZP metabolites
Abbreviations: AADAC, arylacetamide deacetylase; AANZP, 7-acetylamino nitrazepam; ABT, 1-aminobenzotriazole; ANZP, 7-amino nitrazepam; AOX1, aldehyde oxidase 1; BNPP, bis-(p-nitrophenyl) phosphate; CES, carboxylesterase; CM, carboxymethyl; CYP, cytochrome P450; DEAE, diethylaminoethyl; DFP, diisopropyl phosphorofluoride; DMSO, dimethyl sulfoxide; EDTA, ethylenediaminetetraacetic acid; Eserine, sulfate physostigmine; HLC, human liver cytosol; HLM, human liver microsomes; LC–MS/MS, liquid chromatography–tandem mass spectrometry; MNA, N1-methylnicotinamide; MRM, multiple-reaction monitoring; NAC, N-acetyl-L-cysteine; NADP+, nicotinamide adenine dinucleotide phosphate oxidized form; NADPH, nicotinamide adenine dinucleotide phosphate reduced form; NAT, N-acetyltransferase; NZP, nitrazepam; PMSF, phenylmethylsulfonyl fluoride; S9, supernatant 9000g; SDS-PAGE, sodium dodecyl sulfate-poly acrylamide gel electrophoresis; XOR, xanthine oxidoreductase. ⇑ Corresponding author at: Drug Metabolism and Toxicology, Faculty of Pharmaceutical Sciences, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan. E-mail address:
[email protected] (T. Fukami). http://dx.doi.org/10.1016/j.bcp.2017.06.114 0006-2952/Ó 2017 Published by Elsevier Inc.
Please cite this article in press as: K. Konishi et al., Identification of enzymes responsible for nitrazepam metabolism and toxicity in human, Biochem. Pharmacol. (2017), http://dx.doi.org/10.1016/j.bcp.2017.06.114
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Fig. 1. Proposed metabolic pathways and related adverse reactions of NZP.
were detected to a much lesser extent in human urine compared to ANZP and AANZP. The toxicity of NZP has been evaluated by in vitro studies. When HepG2 cells (a hepatocarcinoma cell line) were treated with NZP and recombinant human CYP3A4, significant cytotoxicity was observed compared to the absence of CYP3A4 [6], suggesting that the metabolism NZP is associated with liver injury. However, it remained unclear whether CYP3A4 is the sole enzyme that forms a reactive NZP metabolite. Teratogenicity caused by NZP was reported in rats, but not in mice [7]. ANZP acetylase activity is 8.5-fold higher in rats than in mice, whereas AANZP hydrolase activity is 10-fold lower in rats than in mice. Therefore, AANZP was suggested to be the cause of the observed teratogenicity. As mentioned above, metabolic reactions are thought to be linked with the side effects of NZP, but the enzymes responsible for NZP metabolism in humans are unknown. In rats, NZP has been reported to be reduced by intestinal bacteria [10–12], but it is also conceivable that NZP metabolism efficiently occurs within the liver after being absorbed from the intestinal tract. The identification of the enzymes responsible for NZP metabolism would be helpful to prove the association of NZP metabolism with the pathogenesis of adverse reactions. In this study, we attempted to identify the enzymes responsible for the metabolism of NZP, including reduction, acetylation, and hydrolysis, and to identify the reactive metabolite(s) that may cause liver injuries in humans. 2. Materials and methods 2.1. Materials Pooled human liver cytosol (HLC) (prepared from 150 individuals), pooled human liver microsomes (HLM) (prepared from 50 individuals), pooled human 9000g supernatant (S9) (prepared from 22 individuals), recombinant human CYP1A2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2C19, CYP2D6, CYP2E1, CYP3A4, and CYP3A5 Supersomes, and recombinant N-acetyltransferase (NAT) 1 and NAT2 cytosols were purchased from Corning (Corning, NY). IRDye 680 goat anti-mouse IgG was obtained from LI-COR Biosciences (Lincoln, NE). Acetyl CoA, diisopropyl fluorophosphate (DFP), Nacetyl-L-cysteine (NAC), NZP, phenylmethylsulfonyl fluoride (PMSF), and sulfate physostigmine (eserine) were purchased from Wako Pure Chemicals (Osaka, Japan). Ammonium sulfate, ANZP, and bis(p-nitrophenyl) phosphate (BNPP) were purchased from Sigma-Aldrich (St. Louis, MO). Carboxymethyl (CM) Sepharose
and diethylaminoethyl (DEAE) Sephacel were purchased from GE Healthcare (Buckinghamshine, UK). Glucose-6-phosphate, glucose-6-phosphate dehydrogenase, and b-nicotinamide adenine dinucleotide phosphate (NADP+) were purchased from Oriental Yeast (Tokyo, Japan). 2D-Silver Stain Reagent II and N1methylnicotinamide (MNA) were purchased from Cosmo Bio (Tokyo, Japan). A mouse anti-human aldehyde oxidase 1 (AOX1) monoclonal antibody was purchased from Santa Cruz Biotechnology (Dallas, TX). All other chemicals and solvents were of the highest grade commercially available. 2.2. Measurement of NZP reductase activity NZP reductase activities were determined as follows: a typical reaction (final volume of 200 lL) contained 100 mM potassium phosphate buffer (pH 7.4), enzyme sources (0.4 mg/mL HLM or HLC; 20 lL of a 40–60% ammonium sulfate precipitated fraction and CM Sepharose fraction; or 50 lL of a DEAE Sephacel fraction) and a NADPH-generating system (0.5 mM NADP+, 5 mM glucose6-phosphate, 5 mM MgCl2, and 1 U/mL glucose-6-phosphate dehydrogenase). For the evaluation of the involvement of AOX1, 1 mM MNA was added to the reaction mixture. We confirmed that the ANZP formation rates were linear up to a 1 mg/mL protein concentration and a 90 min (in the absence of MNA) or 30 min (in the presence of MNA) incubation time. NZP was dissolved in dimethyl sulfoxide (DMSO). The reactions were initiated by the addition of NZP at a final concentration of 20 lM after a 2-min preincubation at 37 °C. After a 45- or 10-min incubation (in the absence or presence of MNA, respectively), the reactions were terminated by the addition of 200 lL of ice-cold acetonitrile. After removal of the protein by centrifugation at 20,400g for 5 min, a portion of the supernatant was subjected to liquid chromatography–tandem mass spectrometry (LC–MS/MS). The LC equipment was comprised of a CBM-20A controller (Shimadzu, Kyoto, Japan), LC-20AD pumps (Shimadzu), an SIL-20AC HT autosampler (Shimadzu), a CTO20AC column oven (Shimadzu), and a SPD-20A UV detector (Shimadzu) equipped with a Develosil ODS-UG-3 column (3 lm particle size, 4.6 mm i.d. 150 mm; Nomura Chemical, Seto, Japan). The column temperature was set at 40 °C, and the flow rate was 0.2 mL/min. The mobile phase was 0.1% formic acid/5 mM ammonium formate (A) and acetonitrile containing 0.1% formic acid (B). The conditions for elution were as follows: 15% B (0–2 min), 70% B (2–4 min), 95% B (4–7 min), and 15% B (7–9 min). The LC was connected to a LC-MS 8040 (Shimadzu) and was used in the positive
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electrospray ionization mode. Nitrogen was used as the nebulizing and drying gas at 3 L/min and 15 L/min, respectively. Parent and/or fragment ions were filtered in the first quadrupole and dissociated in the collision cell using argon as the collision gas at 230 kPa. ANZP was monitored in the multiple reaction monitoring (MRM) mode at m/z values of 252.30 and 121.20. The analytical data were processed using LabSolutions (version 5.82 SP1, Shimadzu). 2.3. Purification of the NZP reductase from HLC Human liver samples were supplied by the National Disease Research Interchange (Philadelphia, PA) through the Human and Animal Bridging Research Organization (Chiba, Japan). To purify the enzyme(s) responsible for NZP reduction, HLC was prepared as described previously [13], with all of the procedures carried out at 4 °C. Proteins that were precipitated by the addition of 40–60% saturated ammonium sulfate to HLC were dissolved in 20 mM potassium phosphate buffer (pH 6.5) containing 20% glycerol and 1 mM EDTA, and then were dialyzed against the same buffer to remove the ammonium sulfate. The dialyzed sample was applied to a CM Sepharose cation exchange column (1.25 5 cm) equilibrated with 20 mM potassium phosphate buffer (pH 6.5) containing 20% glycerol and 1 mM EDTA. The proteins were eluted using a gradient method with the equilibration buffer containing 0–1 M KCl at a rate of 0.33 mL/min. The eluate was collected continuously in 3 mL aliquots, and their protein concentrations were determined using the Bradford method [14]. The NZP reductase activities of the fractions were evaluated as described above. Fractions showing activity (fractions 57–61) were pooled and centrifuged at 4250g using Centricon YM-30 concentrators (Millipore, Billerica, MA) to concentrate the proteins and exchange buffer. The concentrated proteins were diluted in 20 mM Tris-HCl buffer (pH 7.4) containing 20% glycerol and 1 mM EDTA and were applied to a DEAE Sephacel anion exchange column (1.25 5 cm) equilibrated with the same buffer. The proteins were eluted using a gradient method with the equilibration buffer containing 0–1 M KCl at a rate of 0.33 mL/min, and the NZP reductase activities of each 3-mL fraction was evaluated. Fractions showing activity (fractions 46–48) were pooled and centrifuged at 6000g using Amicon Ultra-4 10K concentrators (Millipore) to concentrate the proteins. 2.4. SDS-PAGE, silver staining, and Western blotting for AOX1 HLC (23 mg), 40–60% saturated ammonium sulfate (21 mg), pooled CM Sepharose cation exchange chromatography fractions (11 mg), and pooled DEAE Sephacel anion exchange chromatography fractions (0.6 mg) were separated by 10% SDS-PAGE, and the gel was stained using a silver staining kit (Cosmo Bio). To Western blotting for AOX1, each 0.8 mg of HLC, 40–60% saturated ammonium sulfate fraction, pooled CM Sepharose cation exchange chromatography fractions, and pooled DEAE Sephacel anion exchange chromatography fractions were separated by 7.5% SDS-PAGE and transferred to an Immobilon-P transfer membrane (Millipore). The membrane was probed with a mouse anti-human AOX1 primary antibody and a corresponding fluorescent dye-conjugated secondary antibody. The band was detected using an Odyssey Infrared Imaging system (LI-COR Biosciences). 2.5. Inhibition study of NZP reductase activity To evaluate the role of AOX1 in NZP reduction in the human liver, an inhibition study was performed using several inhibitor compounds. Estradiol, estrone, and raloxifene are potent AOX1 inhibitors; ketoconazole, promazine, and simvastatin are moderate AOX1 inhibitors; and diclofenac, metronidazole, and naloxone are weak AOX1 inhibitors [15]. Allopurinol was used as an xanthine
3
oxidoreductase (XOR) inhibitor because the substrate specificity and crystal structure of XOR are close to those of AOX1 [15,16]. It has been reported that allopurinol does not inhibit AOX1 [17]. The final concentrations of all inhibitors were 50 lM. The inhibitors were dissolved in DMSO and the final concentration of DMSO in the reactions was 2%. The experimental procedure and conditions were the same as described above, with 1 mM MNA included in the reaction mixtures The control activity was determined in the presence of 2% DMSO. 2.6. Measurement of ANZP acetylase activity ANZP acetylase activities were determined as follows: a typical reaction (final volume of 200 lL) contained 100 mM potassium phosphate buffer (pH 7.4), enzyme fractions (0.4 mg/mL HLC and 0.025 mg/mL recombinants human NAT1 or NAT2 cytosol), 1 mM acetyl CoA, 100 lM dithiothreitol, and 100 lM EDTA. We confirmed that the AANZP formation rates were linear up to a 1 mg/ mL protein concentration and a 90 min incubation time. ANZP was dissolved in acetonitrile. The reactions were initiated by the addition of ANZP after 2-min preincubation at 37 °C. After a 45min incubation, the reactions were terminated by the addition of 200 lL of ice-cold acetonitrile. After the removal of protein by centrifugation at 20,400g for 5 min, a portion of the supernatant was subjected to LC–MS/MS. The apparatus and conditions were the same as described above. AANZP was detected in the MRM mode at m/z values of 294.05 and 121.15. 2.7. Measurement of AANZP hydrolase activity Because an authentic standard of AANZP is not available, AANZP was produced from ANZP using cytosol containing recombinant NAT2 and was used as a substrate to measure AANZP hydrolase activity. AANZP was biosynthesized as follows: a typical reaction (final volume of 200 lL) contained 100 mM potassium phosphate buffer (pH 7.4), 20 mM ANZP, 0.025 mg/mL NAT2 cytosol, 1 mM acetyl CoA, 100 lM dithiothreitol, and 100 lM EDTA. After a 60 min incubation at 37 °C, ANZP was completely converted to AANZP (data not shown). Reactions were incubated at 100 °C for 3 min to inactivate NAT2, followed by centrifugation at 20,400g for 5 min. AANZP hydrolase activities were measured as follows: an aliquot (150 lL) of the supernatant (corresponding to 15 lM of AANZP in 200 mL of the reaction system) was added to 50 lL of the reaction containing 100 mM potassium phosphate buffer (pH 7.4) and 0.4 mg/mL of the enzyme sources (HLM, Sf21 cell homogenate expressing human carboxylesterase (CES) 1, CES2, and arylacetamide deacetylase (AADAC)) and the reactions (total volume: 200 lL) were incubated for 45 min. The reactions were terminated by the addition of 200 lL of ice-cold acetonitrile. After removal of the protein by centrifugation at 20,400g for 5 min, a portion of the supernatant was subjected to LC–MS/MS and ANZP was measured as described above. 2.8. Inhibition study for AANZP hydrolase activity To evaluate the role of AADAC in AANZP hydrolysis in the human liver, an inhibition study was performed using HLM with standard esterase inhibitors. Organophosphates, such as BNPP and DFP, are general CES inhibitors. PMSF is a general serine esterase inhibitor, but does not inhibit AADAC [18,19]. Eserine and vinblastine are CES2 and AADAC inhibitors [13], and telmisartan is also a CES2 inhibitor [19]. The final concentration of inhibitors used in reactions was 100 lM, except for telmisartan (50 lM). PMSF, vinblastine, and telmisartan were dissolved in DMSO, and the rest were dissolved in distilled water. The final concentration of organic solvents (acetonitrile and DMSO) in the reactions was
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1.75%. The experimental procedure was described above. The control activities were determined in the presence or absence of 1% DMSO.
3. Results
2.9. Detection of reactive metabolite(s) as NAC conjugates and identification of enzymes involved in the reaction(s)
NZP reductase activities in HLC and HLM were measured using a 20 lM substrate concentration (Fig. 2). NZP reductase activity in HLC (0.60 ± 0.04 pmol/min/mg protein) was 6-fold higher than that in HLM (0.10 ± 0.01 pmol/min/mg protein). The NZP reductase activity in HLC increased slightly upon the addition of NADPH (1.5-fold, 0.92 ± 0.03 pmol/min/mg protein). The activity in HLM was also increased by NADPH, but the activity was still lower than that in HLC in the absence of NADPH. These results suggested that reductase(s) other than NADPH-dependent enzymes are responsible for NZP reduction in the human liver.
A previous study suggested that reactive metabolite(s) of NZP appear to be associated with liver injury [6]. NAC is often used to trap reactive metabolites as a nucleophile [20]. To investigate whether the reactive metabolite(s) are produced in the processes of NZP metabolism, we monitored the production of NAC conjugate using LC-MS/MS. A typical reaction (200 mL) was prepared with 100 mM potassium phosphate buffer (pH 7.4), 0.8 mg/mL HLC, and 1 mM MNA. The reactions were initiated by the addition of 100 lM NZP after 2-min preincubation at 37 °C. After a 6-hour incubation, the reactions were terminated by the addition of 200 lL of ice-cold acetonitrile. After the removal of proteins by centrifugation at 20,400g for 5 min, a portion of the supernatants were subjected to LC-MS/MS. To detect the reactive metabolite(s) as NAC conjugate(s), a m/z range of 400–450 was scanned. Since a peak was detected at a m/z 413.05, which corresponds to a NAC conjugate of N-hydroxylamino NZP, a product ion scan was conducted at a m/z range of 100–450 (collision energy: 35 V), setting the m/z 413.05 as the precursor ion, to investigate cleavage pattern of the conjugate. Because the main product ion of the conjugate was 284.05, m/z ion transitions of 413.05 and 284.05 were monitored in MRM mode for the NAC conjugate in subsequent analyses. In general, arylamino compounds were hydroxylated to hydroxylamines by P450s [21]. To investigate whether the NAC conjugate was generated from ANZP via P450s, its formation was monitored using HLM to identify the enzyme(s) responsible for the formation of reactive metabolite(s). The formation of a NAC conjugate from ANZP was evaluated using recombinant human P450 supersomes, and an inhibition study was conducted using HLM. The conjugate formation was determined as follows: a typical reaction (final volume of 200 lL) contained 100 mM potassium phosphate buffer (pH 7.4), enzyme sources (0.4 mg/mL HLM or 10 pmol/mL recombinant human CYP1A2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2C19, CYP2D6, CYP2E1, CYP3A4, or CYP3A5 Supersomes), 5 mM NAC, and 20 mL of an NADPH-generating system. For the inhibition study using HLM, 100 lM a-naphthoflavone [21,22], 10 lM sulfaphenazole [23,24], 100 lM tranylcypromine [24], 10 lM quinidine [23,24], 200 lM erythromycin [25], and 1 mM 1-aminobenzotriazole (ABT) [26] were used as inhibitors of CYP1A2, CYP2C9, CY2C9, CYP2C19, CYP2D6, and CYP3A4/5, respectively, as well as a general P450 inhibitor. All inhibitors were dissolved in acetonitrile. The final concentration of acetonitrile in the incubation mixture was 2%. We confirmed that the NAC conjugate formation rates were linear up to a 0.8 mg/mL protein concentration and a 30 min incubation time. The reactions were initiated by the addition of ANZP after a 2-min (for non-mechanism based inhibitors of P450s) or 30-min (for mechanism based inhibitors of P450 s) preincubation at 37 °C. After a 10-min incubation, the reactions were terminated by the addition of 200 lL of ice-cold acetonitrile. After the removal of protein by centrifugation at 20,400g for 5 min, a portion of the supernatant was subjected to LC–MS/MS and the NAC conjugate was measured as described above.
3.1. NZP reductase activity in the human liver
3.2. Purification and identification of enzymes responsible for NZP reduction in the human liver To identify the enzyme(s) responsible for NZP reduction in the human liver, we purified the enzyme(s) from HLC. HLC was fractionated by the ammonium sulfate precipitation method. The 40–60% ammonium sulfate saturated fraction exhibited the highest reductase activity, and this fraction was subsequently purified using a CM Sepharose cation exchange column. By monitoring the protein concentration and NZP reductase activity of the eluate, we found that fractions 57–61 exhibited NZP reductase activity, with the highest activity in the fraction 57. The pooled proteins (fractions 57–61) were concentrated using a Centricon YM-30 concentrator and subsequently purified using a DEAE Sephacel anion exchange column. The active fractions (fractions 46–48) were pooled and concentrated using an Amicon Ultra-4 10K concentrator, prior to which each fraction was assessed by SDS-PAGE and silver staining, revealing the presence of a number of proteins
2.10. Statistical analysis Statistical significance between two groups was determined by a two-tailed Student’s t test. A value of p < 0.05 was considered statistically significant.
Fig. 2. NZP reductase activity by HLM and HLC in the absence or presence of an NADPH-generating system. HLM and HLC (0.4 mg/mL) were incubated with NZP (20 lM) for 45 min. Each column represents the mean ± S.D. of triplicate determinations. ***P < 0.001 and ###P < 0.001.
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(Fig. 3A). Among them, bands corresponding to 130- and 150-kDa proteins were correlated with the NZP reductase activity (data not shown). The recovery of protein and the NZP reductase activity at each purification step are shown in Table 1. Overall, compared with the initial HLC fraction, we achieved a 24-fold purification of the enzymatically active proteins with a 1.4% yield. The isolated protein was predicted to be AOX1 because this protein has been reported to exhibit bands with 130- and 150-kDa molecular weight by SDS-PAGE [27]. In addition, AOX1 seemed to catalyze the reduction of nitro groups to amino groups, although a limited number of compounds were known as substrates [28]. Our hypothesis was confirmed by Western blotting of a pooled DEAE Sephacel fraction using an anti-human AOX1 antibody (Fig. 3B). 3.3. Kinetic analyses of the NZP reductase activity in HLC To investigate whether AOX1 was responsible for the observed NZP reduction in HLC, kinetic analyses were performed in the presence of MNA, which had been reported to promote AOX1-catalyzed reduction by acting as an electron donor of AOX1 (Table 2 and Fig. 4) [28]. The activities in the absence and presence of MNA fitted the Michaelis-Menten kinetic equation. The Km values of them were similar (25.1 ± 1.0 mM and 21.2 ± 0.4 lM, respectively), but the Vmax value in the presence of MNA was substantially higher (61.8 ± 3.68 pmol/min/mg protein) than that in the absence of MNA (1.33 ± 0.04 pmol/min/mg protein). The CLint value was significantly higher in the presence of MNA (2.92 ± 0.21 lL/min/mg protein) than that in the absence of MNA (0.053 ± 0.003 lL/min/ mg protein). The increase in the NZP reductase activity by MNA supported the role of AOX1 for NZP reduction in the human liver. 3.4. Inhibition studies on NZP reductase activity To further investigate the role of AOX1 in the reduction of NZP in the human liver, inhibition studies were performed using various AOX1 inhibitors (Fig. 5). The activity was completely inhibited
by estradiol, estrone, and raloxifene, which are known as potent inhibitors of AOX1 [15]. The activity was moderately inhibited (% of control, 13.4–27.4%) by ketoconazole, promazine, and simvastatin, but not by diclofenac, metronidazole, naloxone, or allopurinol. Because allopurinol is well known as a potent inhibitor of XOR and exhibits a similar crystal structure and substrate recognition as AOX1 [16], the possibility of the involvement of XOR in NZP reduction could be ruled out. These results suggested that the NZP reduction is primarily catalyzed by AOX1 in the human liver.
3.5. ANZP acetylase activities by HLC, and human recombinant NAT1 and NAT2 It has been reported that N-acetylation of xenobiotics is catalyzed by NAT1 and NAT2 in humans and that these enzymes are highly expressed in the cytoplasm of liver cells [29]. Kinetic analyses of ANZP acetylation were performed using HLC (Fig. 6A). The ANZP could be dissolved in acetonitrile up to 4 mM. Because the acetonitrile concentration in the reaction mixture should be below 1%, the maximum substrate concentration used was 40 lM. The Km and Vmax values for ANZP acetylation could not be calculated because the linearity was observed between the substrate concentration and the reaction rate. The CLint value calculated with the initial slope of the plots of V (velocity) versus S (substrate concentration) was 5.40 ± 0.12 lL/min/mg protein. Next, to identify the isoform responsible for ANZP acetylation, the activities using 20 mM ANZP were measured using recombinant human NAT1 and NAT2. NAT2 showed much higher activity (19.5 ± 0.3 nmol/ min/mg protein) compared to NAT1 (0.043 ± 0.002 nmol/min/mg protein) (Fig. 6B). We confirmed that recombinant NAT1 and NAT2 are functionally active by measuring N-acetylase activities of para-aminosalicylic acid (a substrate of NAT1, 1300 nmol/min/ mg protein at 40 mM) and sulfamethazine (a substrate of NAT2, 470 nmol/min/mg protein at 40 mM), respectively (data not shown). Thus, these results suggest that NAT2 is responsible for ANZP acetylation in the human liver.
Fig. 3. SDS-PAGE, silver staining, and Western blotting for AOX1, in each step of purification fraction. Purification of the enzyme(s) responsible for the NZP reduction from HLC. Protein profiles of HLC fractionated by ammonium sulfate and column chromatography. The proteins were separated by SDS-PAGE and were visualized by (A) silver staining and (B) Western blotting: lane M, molecular weight standard; lane 1, HLC (A: 23 lg of protein; B: 0.8 lg of protein); lane 2, 40–60% ammonium sulfate precipitated fraction (A: 21 lg of protein; B: 0.8 lg of protein); lane 3, CM Sepharose fraction (A: 11 lg of protein; B: 0.8 lg of protein); lane 4, DEAE Sephacel fraction (A: 0.6 lg of protein; B: 0.8 lg of protein). Closed arrowheads indicate 130- and 150-kDa bands correlated with NZP reductase activity.
Table 1 Purification of an enzyme involved in NZP reduction from HLC. Step
Protein content mg
Total activity pmol/min
Specific activity pmol/min/mg
Purification Fold
Recovery %
HLC 40–60% (NH4)2SO4 CM Sepharose DEAE Sephacel
1236 181.5 22.5 0.7
494 1162 119 7
0.4 6.4 5.3 9.5
1 16 13.3 23.8
– 235 24.1 1.4
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Table 2 Kinetic parameters of NZP reductase activity by HLC in the presence or absence of MNA.
MNA ( ) MNA (+)
Km lM
Vmax pmol/min/mg protein
CLint lL/min/mg protein
25.1 ± 1.0 21.2 ± 1.2
1.3 ± 0.0 61.8 ± 3.7***
0.1 ± 0.0 2.9 ± 0.1***
Data are mean ± S.D. of triplicate determinations. *** P < 0.001 compared with absence of MNA.
3.6. AANZP hydrolase activities of HLM, human recombinant CES1, CES2, and AADAC To examine whether AANZP is hydrolyzed in the human liver, HLM containing various expressed hydrolases were assayed. As shown in Fig. 7A, high activity (276 ± 23.7 pmol/min/mg protein at 15 lM AANZP) was observed. The activity was not changed by the addition of NADPH (data not shown). To identify the enzyme responsible for the AANZP hydrolysis, the activity was measured using recombinant human CES1, CES2, and AADAC (Fig. 7A). Among them, only AADAC exhibited activity (546 ± 47.2 pmol/ min/mg protein), indicating that the AANZP is specifically hydrolyzed by AADAC. In our previous study, we confirmed that recombinant CES1, CES2, and AADAC are functionally active by measuring the hydrolase activities of fenofibrate (a substrate of CES1, 84 nmol/min/mg protein at 25 mM), procaine (a substrate of CES2, 4.3 nmol/min/mg protein at 5 mM), indiplon (a substrate of AADAC, 265 pmol/min/mg protein at 1 mM) [30]. 3.7. Inhibition analyses of AANZP hydrolase activity To elucidate the role of AADAC in the hydrolysis of AANZP in the human liver, inhibition studies were performed using various esterase inhibitors (Fig. 7B). The activity in HLM was strongly inhibited only by AADAC inhibitors (i.e., eserine, DFP, vinblastine, and BNPP), whereas it was not inhibited by telmisartan or PMSF [19]. This result supported the hypothesis that AADAC is the enzyme responsible for AANZP hydroxylation in the human liver. 3.8. Detection of reactive metabolite(s)
Fig. 4. Kinetic analysis of NZP reductase activity by HLC. HLC (0.4 mg/mL) was incubated with NZP in the absence and presence of MNA (1 mM) for 45 min and 10 min, respectively. Each point represents the mean ± S.D. of triplicate determinations.
Reactive metabolites are often associated with causing druginduced liver injuries [31]. Arylhydroxylamine is generally considered to be one of the alert structures of drug-induced liver injury because of its high reactivity [32]. Arylhydroxylamine can be produced by the reduction of arylnitro compounds by AOX1 because AOX1 performs this reaction in a two-electron reduction manner [28]. Considering the possibility of N-hydroxylamino NZP production in the process of NZP reduction, we sought to detect Nhydroxylamino NZP by using NAC, which is a trapping agent, to detect a reactive metabolite as a stable conjugate [33]. Because the exact mass of the NAC conjugate derived from hydroxylamino NZP was predicted to be 412, a Q3 scan in the m/z range of 400– 450 was conducted. In the presence of NAC, a peak with a m/z 413.05 was detected with high background noise (data not shown). Therefore, the NAC conjugate was monitored using the single ion monitoring (SIM) mode. Incubation of HLC with NZP in the presence of MNA for 6 hours produced a metabolite with m/z 413.05 (Fig. 8A). Because the spectra also showed a fragment ion at m/z 284.05 by a product ion scan mode, it was conceivable that the metabolite was a NAC conjugate of N-hydroxylamino NZP (Fig. 8B). Beside arylnitro reduction, N-hydroxylamine can be produced via arylamine N-hydroxylation by P450s. When ANZP was incubated with HLM in the presence of an NADPH-generating system and NAC for 6 h, the corresponding conjugate was detected (data not shown). These results indicated that N-hydroxylamino NZP is produced by both metabolic processes of NZP reduction and ANZP N-hydroxylation. 3.9. Identification of enzyme(s) responsible for NAC conjugate formation
Fig. 5. Inhibitory effects of AOX1 and XOR inhibitors on NZP reductase activity in HLC. HLC (0.4 mg/mL) was incubated with NZP (20 lM) for 10 min in the presence of MNA (1 mM) and several inhibitors. Concentration of inhibitors are described in material and methods. The control activity was 25.1 ± 1.2 pmol/min/mg protein. Each column represents the mean ± S.D. of triplicate determinations. N.D.: Not detectable.
To identify the enzyme(s) responsible for ANZP Nhydroxylation, NAC conjugate formation was measured using recombinant human P450s, and inhibition analyses were also performed using HLM. In this study, the conjugate was detected with
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Fig. 6. ANZP acetylase activities by HLC and recombinant human NATs. (A) Kinetic analysis of ANZP acetylase activity by HLC. HLC (0.4 mg/mL) was incubated with NZP for 45 min. Each point represents the mean ± S.D. of triplicate determinations. (B) ANZP acetylase activities by recombinant human NAT1 and NAT2. The cytosol of insect cells (0.025 mg/mL) expressing human NAT1 or NAT2 was incubated with ANZP (20 lM) for 45 min. Each column represents the mean ± S.D. of triplicate determinations. *** P < 0.001 compared to NAT1.
Fig. 7. AANZP hydrolase activities by HLM, recombinant human CES1, CES2, and AADAC. (A) Measurement of AANZP hydrolase activities by HLM, recombinant human CES1, CES2, and AADAC. The cytosol of insect cells (0.025 mg/mL) expressing human NAT2 was incubated with ANZP (20 lM) for 60 min. After incubation, samples were boiled for 3 min, followed by centrifugation. A hundred fifty mL of the supernatant, HLM (0.4 mg/mL), and recombinant esterases (0.4 mg/mL), were mixed and incubated for 45 min. Each column represents the mean ± S.D. of triplicate determinations. N.D.: Not detected. (B) Inhibitory effects of serine hydrolase inhibitors on AANZP hydrolase activity in HLM. Concentrations of HLM were 0.4 mg/mL and inhibitors were 100 lM except for telmisartan (50 lM), respectively. The control activity was 387 ± 66.9 pmol/min/mg protein. Each column represents the mean ± S.D. of triplicate determinations.
MRM mode (m/z 413.05 > 284.05) using LC-MS/MS. Among various isoforms, the conjugate was highly formed in the presence of recombinant CYP3A4 and CYP3A5, followed by CYP1A2, CYP2D6, CYP2C19, and CYP2B6 (Fig. 9A). The conjugate formation in HLM was significantly inhibited by ABT (a general P450 inhibitor) and erythromycin (a CYP3A4/5 inhibitor), whereas it was enhanced by a-naphthoflavone (a hetero-activator of CYP3A4) [34]. The extent of inhibition by the other chemicals was not prominent. These results suggested that ANZP is primarily hydroxylated by CYP3A4 in the human liver. 4. Discussion NZP, which is widely used as a hypnotic agent, rarely causes liver injury in humans and teratogenicity in rodents [4–7]. These toxicities may be associated with metabolites of NZP that have been demonstrated to be produced in in vitro or in vivo studies
[6,7]. However, the metabolic enzymes that have been suggested to cause these adverse reactions have not been identified. In this study, we sought to identify enzymes involved in NZP metabolism in humans to elucidate the relationship between metabolism and adverse NZP reactions. We first found that NZP reductase activity in HLC was 6-fold higher than that in HLM (Fig. 2). Considering the fact that the protein content in the cytosol is approximately 5-fold higher than that in microsomes [35], it is clear that the reductase(s) in HLC were primarily involved in the NZP reduction. It should be noted that NADPH did not greatly affect the NZP reductase activities in HLC and HLM (Fig. 2). Although reductases involved in drug metabolism, such as aldo-keto reductases, carbonyl reductases, and NADPH-quinone oxidoreductases, are localized in the liver cytosol [36–38], their contribution would be low because they require NADPH to exert their activities. In addition, microsomal enzymes catalyzing drug reduction, including P450, NADPH-P450
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Fig. 8. Mass spectra of NAC conjugate. (A) Mass spectra with SIM (m/z 413.05) mode in LC–MS/MS analysis of NAC conjugate. HLC (0.8 mg/mL) was incubated with NZP (100 lM) and 5 mM NAC in the presence of 1 mM MNA for 6 h. (B) Mass spectra with product ion scan mode (m/z 413.05 > 100–450) and cleavage pattern of NAC conjugate formed in the reaction described above.
Fig. 9. A NAC conjugate formation from ANZP in the presence of recombinant P450s and HLM. (A) Measurement of a NAC conjugate formation from ANZP in the presence of recombinant human P450s supersomes. Human CYP1A2, CYP2A6, CYP2B6, CYP2C8, CYP2C9, CYP2C19, CYP2D6, CYP2E1, CYP3A4, and CYP3A5 Supersomes (2 pmol P450s) were incubated with ANZP (20 lM) in the presence of an NADPH-generating system and 5 mM NAC for 10 min. Each column represents the mean ± S.D. of triplicate determinations. N.D.: Not detected. A NAC conjugate formation was not detected by Supersomes insect cell microsomes, negative control (Corning, data not shown). (B) Inhibitory effects of several P450s inhibitors on the activity in HLM. HLM (0.4 mg/mL) were incubated with ANZP (20 lM) in the presence of NADPH for 10 min in the presence of NADPH-generating system and 5 mM NAC. Each column represents the mean ± S.D. of triplicate determinations.
oxidoreductase, and cytochrome b5 also hardly contribute to the NZP reduction [39–41]. To date, NADPH-independent nitro-reductases involved in drug metabolism are not well understood. Therefore, we sought to pur-
ify the NADPH-independent NZP reductase(s) from HLC, and identified AOX1 (Fig. 3). We predicted that specific activity in DEAE Sephacel fraction should be more increased compared with the previous one (Table 1). It has been reported that AOX1 protein is
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unstable [42], therefore AOX1 protein may be gradually decomposed or inactivated during the purification process. AOX1 is a molybdo-flavoenzyme that is known as a drug-metabolizing enzyme, catalyzing the oxidation of N-heterocyclic drugs such as famciclovir and zaleplon [43]. In addition, it has been recently reported that AOX1 catalyzes amide-hydrolysis of GDC-0834, which is a Bruton’s tyrosine kinase inhibitor being investigated as a therapeutic agent for rheumatoid arthritis [44]. Thus, AOX1 appears to be able to catalyze various Phase I reactions. To our knowledge, there are no reports that clearly elucidate the role of AOX1 in the nitro-reduction of therapeutic drugs, although rabbit AOX has been shown to catalyze nitro-reduction of 1-nitropyrene and 3-nitrofluoranthene, which are environmental pollutants [28]. Bauer and Howard [28] showed that the AOX1 reductase activity was increased by MNA, which is a metabolite of niacin in the liver, by acting as an electron donor to AOX1. Similarly, in our study, the NZP reductase activity in HLC was enhanced in the presence of MNA (Fig. 4). This is the first study to show that AOX1 catalyzes nitro reduction of a clinically used drug. The role of AOX1 in NZP reduction was supported by an inhibition study (Fig. 5). Although we tried to evaluate the reductase activity using recombinant AOX1 expressed in Escherichia coli (Cypex, UK), the activity was scarcely detected (data not shown). We also measured the marker activity of AOX1, phthalazine oxidase activity, but the activity by recombinant AOX1 (82.2 pmol/min/mg at 100 mM) was much lower than that in HLC (1660.4 pmol/min/mg at 100 mM). Even if the NZP reduction was occurred with 20-fold lower efficiency compared with that by HLC, the peak in LC-MS/ MS could not be appeared because of the detection limit. This recombinant was the only commercially available one. Therefore, the NZP reductase activity could not be evaluated using recombinant AOX1. Human XOR, which is also a molybdo-flavoenzyme, shares a 40% amino acid identity with the human AOX1. XOR has similar molecular structure and substrate recognition properties with AOX1 [16]. However, inhibition studies did not support the involvement of XOR in NZP reduction. Thus, we could provide important information with regards to a novel function of AOX1 as a drug-metabolizing enzyme. The N-acetylation of clinical drugs is catalyzed by NAT1 and NAT2 in humans [45]. For ANZP, recombinant human NAT2 showed significantly higher acetylase activity than NAT1, clearly indicating that NAT2 is responsible for ANZP acetylation (Fig. 6B). There is a large interindividual variation of NAT2 enzymatic activity in humans owing to genetic polymorphisms [46]. In fact, it has been reported that the AANZP/ANZP ratios in urine 8–10 h after
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oral administration of NZP exhibited a 4.5-fold interindividual variation [8]. Both NAT2⁄5B (c.341T > C/481C > T/803A > G) and NAT2⁄6A (c.282C > T/590G > A) are predominant variants that result in a significant decrease of NAT2 enzyme activity [45]. Clinical studies needs to be conducted to elucidate whether subjects with these variants show low plasma AANZP and high ANZP levels. A previous report indicated that AANZP is hydrolyzed in the rodent liver [7]. We found that the hydrolysis also occurs in HLM (Fig. 7A). The hydrolysis was also observed in HLC, but its activity was approximately 44-fold lower than in HLM (data not shown). Analyses using recombinant enzymes in an inhibition study clearly demonstrated that AADAC, which is localized in microsomes, is responsible for AANZP hydrolysis in the human liver. The activity observed in HLC may be due to a contamination of AADAC in HLC. We recently clarified that AADAC prefers small acyl substrate moieties [47]. Because AANZP has a small acyl moiety, the role of AADAC in AANZP hydrolysis was plausible. Takeno et al [7] have reported that AANZP hydrolase activity in mouse liver microsomes is approximately 10-fold higher than rat liver microsomes [7]. We have previously found that AADAC mRNA expression level in the mouse liver is 10-fold higher than that in the rat liver, and AADAC-catalyzed phenacetin hydrolysis in mouse liver microsomes was 6.5-fold higher than in rat liver microsomes [13]. Taken together, AADAC is likely responsible for AANZP hydrolysis in rodents as well. Our previous report showed a 22 to 205-fold interindividual variation in the activity of AADAC in humans [48]. This variation has been partly explained by genetic polymorphisms, especially AADAC⁄3 (g.13651G > A/g.14008T > C), which causes a substantial decrease in AADAC enzyme activity [49]. In addition to NAT2, AADAC could also cause interindividual variation in the pharmacokinetics of NZP. Drug-induced liver injury is a critical problem in drug development [30]. In general, arylhydroxylamine is thought to be linked with hepatotoxicity because of its high reactivity. For example, flutamide, which is used for the treatment of prostate cancer, is ultimately metabolized to a hydroxylamine that is suggested to cause hepatotoxicity in vivo [50]. Dapsone, which is used for the treatment of leprosy, has been suggested to be metabolically converted to a hydroxylamine that may covalently bind to protein, leading to oxidative stress and hepatotoxicity [51,52]. Flutamide and dapsone have a primary amine structure and are N-hydroxylated to hydroxylamine by P450 s. In addition to amine hydroxylation, hydroxylamine can be formed by nitro reduction [28]. In this study, we found that both steps for the reduction of NZP and the hydroxylation of ANZP produce the hydroxylamine derivative of NZP (Figs. 8
Fig. 10. Clarified metabolic pathways and responsible enzymes involved in NZP metabolism in human.
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and 9). Because NAT2 catalyzes the acetylation of ANZP, it may also regulate the level of ANZP in the liver. Thus, the N-hydroxylamino NZP level in the liver would be determined by multiple metabolic enzymes, including AOX1, AADAC, CYP3A4, and NAT2. We previously reported that the cytotoxicity of NZP in HepG2 cells was enhanced by an incubation with recombinant human CYP3A4 Supersomes, but not with CYP2C9 and CYP2C19 Supersomes [6]. In that study, we considered that the increased cytotoxicity could be due to the increase in N-hydroxylation of AZNP by CYP3A4 (Fig. 9). However, in HepG2 cells, the AOX1 protein was not detected by Western blotting (data not shown). In addition, the NAC conjugate was not detected when NZP was incubated with HLM in the presence of NADPH (data not shown). Therefore, the cytotoxicity of NZP observed in the previous report [6] might be due to the reactive metabolite produced by ANZP N-hydroxylation. In conclusion, we have elucidated the enzymes responsible for NZP metabolism in the human liver (Fig. 10). There are large interindividual variabilities in the activities of AOX1, NAT2, AADAC, and CYP3A4 [44,53–55] that could affect the sensitivity of individuals to the adverse reactions of NZP. The present study provides basic information needed to consider the pharmacokinetics and adverse reactions of NZP in humans. Since the present study does not show the direct evidence for the mechanism of the NZPinduced liver injury, in vitro or in vivo studies to directly prove the association of metabolic pathways with NZP toxicity are required in the near future. Conflict of interest The authors declare no conflict of interest. Author contributions 1. Study conception and design: KK, TF; 2. Acquisition, analysis and/or interpretation of data: KK, TF, SG, MN; 3. Drafting/revision of the work for intellectual content and context: KK, TF, MN; 4. Final approval and overall responsibility for the published work: KK, TF, SG, MN Acknowledgement This work was supported in part by a Grant-in-Aid for Scientific Research (C) from the Japan Society for the Promotion of Science (16K08367). References [1] E. Fortier-Brochu, S. Beaulieu-Bonneau, H. Ivers, C. Morin, Insomnia and daytime cognitive performance: a meta-analysis, Sleep Med. Rev. 16 (2012) 83–94. [2] E. Ford, T. Cunningham, W. Giles, J. Croft, Trends in insomnia and excessive daytime sleepiness among US adults from 2002 to 2012, Sleep Med. 16 (2015) 372–378. [3] A. Holbrook, R. Crowther, A. Lotter, The role of benzodiazepines in the treatment of insomnia, J. Am. Geriatr. Soc. 49 (2001) 824–826. [4] R. Andrade, J. Agundez, M. Lucena, C. Martinez, R. Cueto, E. Garcia-Martin, Pharmacogenomics in drug induced liver injury, Curr. Drug Metab. 10 (2009) 956–970. [5] X. Zhu, N. Kruhlak, Construction and analysis of a human hepatotoxicity database suitable for QSAR modeling using post-market safety data, Toxicology 321 (2014) 62–72. [6] K. Mizuno, M. Katoh, H. Okumura, N. Nakagawa, T. Negishi, T. Hashizume, M. Nakajima, T. Yokoi, Metabolic activation of benzodiazepines by CYP3A4, Drug Metab. Dispos. 37 (2009) 345–351. [7] S. Takeno, Y. Hirano, A. Kitamura, T. Sakai, Comparative developmental toxicity and metabolism of nitrazepam in rats and mice, Toxicol. Appl. Pharm. 121 (1993) 233–238. [8] A. Karim, D. Evans, Polymorphic acetylation of nitrazepam, J. Med. Genet. 13 (1976) 17–19. [9] H. Sawada, K. Shinohara, On the urinary excretion of nitrazepam and its metabolites, Arch. Toxicol. 28 (1971) 214–221.
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Please cite this article in press as: K. Konishi et al., Identification of enzymes responsible for nitrazepam metabolism and toxicity in human, Biochem. Pharmacol. (2017), http://dx.doi.org/10.1016/j.bcp.2017.06.114