Aromatic hydroxylation and catechol formation: A novel metabolic pathway of the growth promotor zeranol

Toxicology Letters 192 (2010) 379–386

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Aromatic hydroxylation and catechol formation: A novel metabolic pathway of the growth promotor zeranol Andreas Hildebrand, Erika Pfeiffer, Manfred Metzler ∗ Institute of Applied Biosciences, Chair of Food Chemistry, Karlsruhe Institute of Technology (KIT), Karlsruhe, Germany

a r t i c l e

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Article history: Received 15 September 2009 Received in revised form 7 November 2009 Accepted 11 November 2009 Available online 18 November 2009 Keywords: Zeranol ␣-Zearalanol Zearalanone Aromatic hydroxylation Catechols Growth promotor

a b s t r a c t ␣-Zearalanol (␣-ZAL, zeranol) is a macrocyclic resorcylic acid lactone, which is highly estrogenic and used as a growth promotor for cattle in various countries. Little is known about the phase I metabolism of ␣-ZAL. We now report that ␣-ZAL and its major metabolite zearalanone (ZAN) are extensively monohydroxylated at the aromatic ring by microsomes from human liver in vitro. This novel pathway leads to catechols, the chemical structures of which were unambiguously established by the use of deuterium-labeled ␣-ZAL and ZAN, and by the synthesis of authentic standards. The aromatic hydroxylation of ␣-ZAL is almost exclusively mediated by the human cytochrome P450 (hCYP) 1A2 isoform. The catechol metabolites of ␣-ZAL and ZAN are unstable and readily oxidized to quinones, which could be detected among the metabolites of ␣-ZAL and ZAN generated by human hepatic microsomes and hCYP1A2. Furthermore, the quinone metabolites are able to form covalent adducts with N-acetylcysteine (NAC), as several of such adducts were found in microsomal incubations fortified with NAC. Aromatic hydroxylation of ␣ZAL was also observed with bovine, porcine and rat hepatic microsomes. Further studies are needed to demonstrate the catechol pathway of ␣-ZAL in vivo and to assess its toxicological significance. © 2009 Elsevier Ireland Ltd. All rights reserved.

1. Introduction ␣-Zearalanol (␣-ZAL, generic name zeranol, Fig. 1) is a macrocylic resorcylic acid lactone with a pronounced estrogenic activity (Lindsay, 1985; Takemura et al., 2007). It is prepared industrially by the reduction of the estrogenic mycotoxin zearalenone (ZEN, Fig. 1), which is produced by fungi of the species Fusarium and frequently contaminates corn and other crops (Bennett and Klich, 2003). Due to its high hormonal activity, ␣-ZAL is legally used in the USA, Canada and several other countries, but not in the European Union, as a growth promotor in cattle under the trade name RalGro® (Wang and Wang, 2007). The metabolism of ␣-ZAL has been studied in several species including humans, and both phase I and phase

Abbreviations: COMT, catechol-O-methyltransferase; CYP, cytochrome P450; DAD, diode array detector; E1, estrone; E2, 17␤-estradiol; ESI, electrospray ionization; GC, gas chromatography; hCYP, human CYP; LC, liquid chromatography; MS, mass spectrometry; MSn , daughter ion mass spectrometry; NAC, N-acetylcysteine; NAD(H), nicotinamide adenine dinucleotide (reduced form); NADP(H), nicotinamide adenine dinucleotide phosphate (reduced form); Rt, retention time; t-BME, tert-butyl methyl ether; UV, ultraviolet; ZAL, zearalanol; ZAN, zearalanone; ZEN, zearalenone. ∗ Corresponding author at: Institute of Applied Biosciences, Karlsruhe Institute of Technology (KIT), Bldg. 50.41, Adenauerring 20a, 78131 Karlsruhe, Germany. Tel.: +49 721 608 2132; fax: +49 721 608 7255. E-mail address: [email protected] (M. Metzler). 0378-4274/$ – see front matter © 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.toxlet.2009.11.014

II metabolites have been identified (Baldwin et al., 1983; Migdalof et al., 1983). The major phase I metabolite was zearalanone (ZAN, Fig. 1), arising through dehydrogenation of the aliphatic hydroxy group at C-7, whereas a minor metabolite was ␤-ZAL (Fig. 1, also called taleranol), which is the stereoisomer of ␣-ZAL and probably formed by reduction of ZAN. Phase II metabolites comprise both glucuronides and sulfates of ␣-ZAL and ZAN. Thus, the metabolism of ␣-ZAL is very similar to that reported for the mycotoxin ZEN so far, which is also characterized by reduction/oxidation at C-7 and formation of glucuronide and sulfate conjugates (Zinedine et al., 2007). Our laboratory has recently disclosed that ZEN is not only metabolized at the aliphatic macrocycle but also at the aromatic ring: hydroxylation at C-15 and, to a lesser extent, at C-13, were found to constitute major metabolic pathways of ZEN in vitro, especially with microsomes from human liver (Pfeiffer et al., 2009). Therefore, the objective of the present study was to probe ␣-ZAL for aromatic hydroxylation. We report here that the catechols 13hydroxy-␣-ZAL and 15-hydroxy-␣-ZAL are major metabolites of ␣-ZAL with hepatic microsomes from humans, and are also generated by rat, bovine and porcine liver microsomes. We have now unequivocally established the structures of these novel metabolites and elucidated the human cytochrome P450 (hCYP) isoforms catalyzing the aromatic hydroxylation of ␣-ZAL. Moreover, preliminary evidence was obtained that the catechol metabolites are easily oxidized to quinones capable of covalently binding to thiols. Part of

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A. Hildebrand et al. / Toxicology Letters 192 (2010) 379–386 48 h. The crude product, consisting of 25% starting material and 75% 13,15-D2 -ZAN, was used for the next step without purification. 6,6,8,8,13,15-D6 -␣-ZAL: 5 mg NaBH4 was added to the solution of 0.05 mmol 6,6,8,8,13,15-D6 -ZAN in 0.5 ml MeOH and the mixture magnetically stirred at 20 ◦ C for 1 h. The solvent was removed under reduced pressure, 3 ml water was added and the aqueous phase extracted with 2× 5 ml t-BME. The ␣- and ␤-stereoisomers of 6,6,8,8,13,15-D6 -ZAL were separated by preparative HPLC. 13,15-D2 -␣-ZAL: The crude 13,15-D2 -ZAN (about 0.04 mmol) was reduced with NaBH4 and the ␣- and ␤-stereoisomers of 13,15-D2 -ZAL were separated by preparative HPLC as described for 6,6,8,8,13,15-D6 -ZAL.

Fig. 1. Chemical structures of ␣-ZAL, ␤-ZAL, ZAN and ZEN.

this work has been presented at the 2009 Meeting of the German Society of Pharmacology and Toxicology (Hildebrand et al., 2009). 2. Materials and methods

2.2.2. 13-Hydroxy-˛-ZAL and 15-hydroxy-˛-ZAL These compounds were accessible from 13-nitro-ZEN and 15-nitro-ZEN, prepared by nitration of ZEN and separation of the resulting mixture of mononitro-ZEN isomers by preparative HPLC as reported earlier (Pfeiffer et al., 2009). Catalytic hydrogenation of the pure nitro-ZEN isomers afforded the respective amino-ZANs, which were subsequently autoxidized to the quinone imines, hydrolyzed to the quinones, and reduced to the catechols (Fig. 3). Conversion of 13-hydroxy-ZAN and 15-hydroxy-ZAN to a mixture of the ␣- and ␤-stereoisomers of 13-hydroxy-ZAL and 15-hydroxy-ZAL was achieved with NaBH4 . 13-Amino-ZAN and 15-amino-ZAN: A solution of 0.01 mmol 13-nitro-ZEN or 15-nitro-ZEN in 1 ml ethyl acetate containing 1 mg 10% Pd/C was magnetically stirred at 20 ◦ C under a hydrogen atmosphere for 15 h, after which the catalyst was removed by centrifugation. LC–DAD–MS analysis showed that the respective amino-ZAN was formed in quantitative yield with a purity of >99%. 13-Amino-ZAN: LC–MS data (Rt 9.0 min): M−H ion at m/z 334; MS2 for 334: 334 (2), 316 (17), 306 (4), 290 (100), 272 (5).

2.1. Purchased chemicals ZEN was purchased from Fermentek (Jerusalem, Israel), ZAN and the ␣- and ␤isomers of ZAL from Sigma/Aldrich/Fluka (Taufkirchen, Germany). All compounds had a purity of >98% according to HPLC–DAD analysis. Chemicals used for the synthesis of reference compounds, nicotinamide adenine dinucleotide (NAD+ ) and its 5 -phosphate (NADP+ ), and other chemicals and reagents were of the highest quality available and were also purchased from Sigma/Aldrich/Fluka. HPLC grade acetonitrile was from Acros Organics (Geel, Belgium). 2.2. Synthesized chemicals 2.2.1. Deuterium-labeled ZAN and ˛-ZAL Two types of deuterium-labeled ␣-ZAL were prepared from ZEN (Fig. 2): hexadeutero-␣-ZAL was labeled at the aliphatic positions 6 and 8 next to the carbonyl group as well as at the aromatic positions 13 and 15, whereas dideutero␣-ZAL was specifically labeled at the aromatic positions 13 and 15. In each case, the respective ZANs were synthesized first and subsequently reduced with sodium borohydride to a mixture of ␣- and ␤-ZAL, which were separated by preparative HPLC using the conditions described under Section 2.5. 6,6,8,8,13,15-D6 -ZAN was prepared from ZAN by alkaline deuterium exchange in deuterated water, using a modification of the method reported earlier for the deuterium-labeling of ZEN (Miles et al., 1996). 13,15-D2 -ZAN was obtained by the same exchange reaction from ZAN-7-[1,3]dioxolane and subsequent hydrolysis of the dioxolane. Analysis by LC–MS in the negative ESI mode gave the following relative intensities of the M−H ions: for 6,6,8,8,13,15-D6 -␣-ZAL: m/z 327:326:325:324 = 100:68:20:3; for 13,15D2 -␣-ZAL: m/z 323:322:321 = 100:15:2, indicating a near quantitative exchange of hydrogen. ZAN: 0.5 mmol (159 mg) ZEN in 10 ml methanol was magnetically stirred under a hydrogen atmosphere in the presence of 20 mg of a 10% Pd on charcoal catalyst at 20 ◦ C for 15 h. The catalyst was removed by centrifugation, washed with ethyl acetate to dissolve precipitated ZAN, and the combined supernatant concentrated to dryness using a rotary evaporator. The product was >99.9% pure ZAN as identified by its LC–MS retention time and ESI-MS in comparison with authentic ZAN. ZAN-7-[1,3]dioxolane: 0.05 mmol ZEN-7-[1,3]dioxolane was prepared according to Cramer et al. (2007) from ZEN and ethylene glycol. The crude product still containing 15% ZEN was dissolved in 1.0 ml methanol and hydrogenated as described for ZAN. The product was free of ZEN and ZEN-dioxolane according to LC–DAD–MS, and was directly used for the deuterium-labeling. 6,6,8,8,13,15-D6 -ZAN: 0.05 mmol ZAN dissolved in 200 ␮l tetrahydrofurane was added to the solution of 125 mg K2 CO3 in 1.0 ml D2 O. After 144 h at 20 ◦ C in the dark, 3 ml of water was added and the mixture neutralized to pH 7 with 1 M aqueous HCl, followed by extraction with 10 and 5 ml t-BME. The organic phase was dried over MgSO4 and contained the hexadeuterated ZAN with a purity of >99% according to LC–DAD–MS. 13,15-D2 -ZAN-7-[1,3]dioxolane: 0.05 mmol crude ZAN-7-[1,3]dioxolane was subjected to deuterium exchange as described for ZAN. The crude product contained about 85% of the desired product and was purified by preparative HPLC. 13,15-D2 -ZAN: 0.05 mmol 13,15-D2 -ZAN-7-[1,3]dioxolane, dissolved in 1.5 ml MeOH and 0.5 ml 1 M aqueous NH4 Cl solution, was magnetically stirred at 50 ◦ C for

Fig. 2. Chemical synthesis of two different deuterium-labeled ZANs and ␣-ZALs. (a) Ethylene glycol/H+ and (b) H2 /Pd/C.

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LC–DAD–MS analysis, 13-hydroxy-ZAN was formed in high yield and with a purity of 90%, whereas the yield of 15-hydroxy-ZAN was only about 5%, probably due to the lower chemical stability of this isomer. 13-Hydroxy-ZAN: LC–MS data (Rt 18.6 min): M−H ion at m/z 335; MS2 for 335: 317 (55), 307 (13), 291 (100), 273 (13), 221 (12). 15-Hydroxy-ZAN: LC–MS data (Rt 18.0 min): M−H ion at m/z 335; MS2 for 335: 317 (100), 307 (2), 291 (46), 273 (7), 221 (8). 13-Hydroxy-␣- and ␤-ZAL and 15-hydroxy-␣- and ␤-ZAL: 2 mg NaBH4 was added to the solution of the respective hydroxy-ZAN (about 0.01 mmol) in methanol. After magnetic stirring at 20 ◦ C for 30 min, the solution contained a mixture of the ␣and ␤-isomers of the respective hydroxy-ZAL, which were separated by preparative HPLC.

13-Hydroxy-˛-ZAL: LC–MS data (Rt 15.0 min): M−H 319 (49), 309 (6), 293 (100), 275 (13). 13-Hydroxy-ˇ-ZAL: LC–MS data (Rt 12.6 min): M−H 319 (49), 309 (6), 293 (100), 275 (12). 15-Hydroxy-˛-ZAL: LC–MS data (Rt 14.5 min): M−H 319 (100), 309 (4), 301 (11), 293 (81), 275 (20). 15-Hydroxy-ˇ-ZAL: LC–MS data (Rt 13.2 min): M−H 319 (100), 309 (3), 301 (11), 293 (82), 275 (19).

ion at m/z 337; MS2 for 337: ion at m/z 337; MS2 for 337: ion at m/z 337; MS2 for 337: ion at m/z 337; MS2 for 337:

NAC adducts of 13-hydroxy-ZAN and 15-hydroxy-ZAN: A solution of 1 ␮mol of the synthetic catechol in 10 ␮l DMSO was mixed with 375 ␮l of 0.1 M potassium phosphate buffer pH 7.4 and 125 ␮l of a 40 mM aqueous NAC solution. After shaking for 90 min at 20 ◦ C in the presence of air, 500 ␮l of 0.7 M glycine/HCl buffer pH 1.2 was added and the NAC adducts were extracted with 2× 500 ␮l ethyl acetate.

2.3. Cell fractions Pooled human hepatic microsomes from 24 donors (cat. no. 452161, lot no. 18888, 12 males and 12 females; 23 Caucasians and 1 Hispanic, with ages ranging from 33 to 78 years) were purchased from BD Gentest (Woburn, MA, USA). Microsomes were prepared from the fresh livers of male Wister rats, steers and female pigs from the local slaughterhouse in our laboratory according to the method of Lake (1987). The CYP content, determined according to Omura and Sato (1964), of human, rat, bovine and porcine microsomes was 0.30, 0.67, 0.69 and 0.47 nmol/mg protein, respectively. Supersomes, i.e. microsomes from insect Sf-9 cells infected with a baculovirus strain containing the cDNA of human CYP1A1, 1A2, 1B1, 2A6, 2B6, 2C8, 2C9, 2C19, 2D6 or 3A4 were also from BD Gentest. The activity of the supersomes was certified by the supplier. Moreover, the same supersomes were used in a recent study on the hydroxylation of four Alternaria toxins, for which many of the CYP isoforms were found to exhibit high activities (Pfeiffer et al., 2008).

2.4. Incubations with hepatic microsomes and CYP isoforms

Fig. 3. Synthesis of the ␣- and ␤-isomers of 13-hydroxy-ZAL and 15-hydroxy-ZAL. The chemical structure of the quinone imines and quinones is tentative, and isomeric structures are possible. 15-Amino-ZAN: LC–MS data (Rt 10.6 min): M−H ion at m/z 334; MS2 for 334: 334 (1), 316 (100), 290 (44), 272 (4), 220 (10). 13-Hydroxy-ZAN and 15-hydroxy-ZAN: The solution of the respective amine (about 0.01 mmol) in ethyl acetate was concentrated to dryness under a stream of nitrogen, and the residue dissolved in 200 ␮l methanol. 100 ␮l of 10% aqueous K2 CO3 was added and the mixture stirred magnetically at 20 ◦ C in the presence of air. Within a few minutes, a brownish color developed, which turned yellow after adding 500 ␮l of diluted sulfuric acid (100 ␮l conc. H2 SO4 plus 400 ␮l water). The yellow material was extracted with 500 ␮l t-BME, which was subsequently removed and added to a magnetically stirred solution of 20 mg (95 ␮mol) of Na2 S2 O4 × 2H2 O in 1 ml of water. After 10 min, the now slightly yellow t-BME phase was separated, washed with 200 ␮l of saturated aqueous NaCl, and dried over Na2 SO4 . According to

Incubation with hepatic microsomes was carried out in a total volume of 500 ␮l of 0.1 M potassium phosphate buffer pH 7.4. The substrate concentration was 100 ␮M and the microsomal concentration 1 mg protein per ml. After 5 min preincubation at 37 ◦ C, the enzymatic reaction was started by adding 17.5 ␮l of a NADPH-generating system, containing 625 ␮g NADP+ , 2.5 ␮mol MgCl2 , 5.2 ␮mol isocitrate and 0.25 U isocitrate dehydrogenase, and kept at 37 ◦ C for 40 min. The mixture was then put on ice and extracted twice with 500 ␮l ethyl acetate each. The combined extract was evaporated to dryness under a stream of nitrogen, and the residue dissolved in 50 ␮l methanol for LC–DAD–MS analysis. Control incubations were carried out in the absence of the NADPH-generating system. Individual hCYP isoforms were delivered as suspension in 500 ␮l buffer with a concentration of 1 nmol CYP per ml. The assays for measuring their enzymatic activities were conducted according to the protocol provided by BD Gentest, taking into account the suitable incubation buffer and incubation time for each isoform to ensure linear kinetics of product formation. In a typical assay with a recombinant hCYP isoform, 5 ␮l of a 10 mM solution of ␣-ZAL in DMSO and 5 ␮l of the suspension of the hCYP isoform (concentration 1 pmol/␮l) were added to 500 ␮l of 0.1 M potassium phosphate buffer pH 7.4 or 0.05 M tris/chloride buffer pH 7.5. After preincubation at 37 ◦ C for 1 min, 17.5 ␮l of the NADPH-generating system was added and incubations continued for 10 min. Workup was as described for incubations with mammalian microsomes, and extracted metabolites were analyzed by analytical HPLC. Control incubations were conducted with microsomes devoid of a hCYP isoform. For the formation of NAC adducts, microsomal or supersomal incubations were conducted as described above but only in 375 ␮l buffer for 10 min. 125 ␮l of 40 mM aqueous NAC solution was then added and incubations continued at 37 ◦ C for another 30 min. Extraction was carried out as described for the NAC adducts of the synthetic catechols (see Section 2.2.2).

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Table 1 Characterization of ␣-ZAL metabolites generated by NADPH-fortified human hepatic microsomes and analyzed by LC–DAD–MS. Compounda

Rt LC–MS (min)

ESI-MSb (M−H)

ESI-MS2 of M−H (relative intensity)

␣-ZAL ␤-ZAL ZAN Peak 1 Peak 2 Peak 3 Peak 4 Peak 5 Peak 6 Peak 7 Peak 8

18.0 16.1 21.7 13.3 14.5 14.8 15.0 16.4 17.5 18.0 18.6

321 321 319 337 337 335 337 335 333 335 335

303 (35), 293 (4), 277 (100), 259 (5) 303 (35), 293 (4), 277 (100), 259 (5) 301 (25), 277 (4), 275 (100), 205 (19), 163 (6) 319 (57), 293 (100), 275 (25), 221 (32), 177 (19) 319 (100), 309 (4), 301 (11), 293 (79), 275 (17) 335 (1), 317 (12), 307 (100), 291 (67), 263 (12) 319 (45), 309 (6), 293 (100), 275 (12) 317 (100), 291 (85), 273 (26), 247 (4), 161 (9) 315 (15), 305 (100), 289 (67), 271 (8), 261 (6) 317 (100), 307 (3), 291 (43), 273 (6), 221 (8) 317 (53), 307 (13), 291 (100), 273 (13), 221 (11)

a b

Mass shift of M−H for D6 -labeled

D2 -labeled

+6 +6 +6 +5 +5 +4 +5 +5 +4 +5 +5

+2 +2 +2 +2 +1 0 +1 +2 0 +1 +1

Chemical structure

␣-ZAL ␤-ZAL ZAN 6- or 8-HO-␣-ZAL 15-HO-␣-ZAL 13-HO-␣-ZAL-quinone 13-HO-␣-ZAL 6- or 8-HO-ZAN 13-HO-ZAN-quinone 15-HO-ZAN 13-HO-ZAN

Designation as in Fig. 4. Negative mode.

2.5. Analytical and preparative HPLC For analytical HPLC, a Hewlett Packard 1100 system equipped with a binary pump, a DAD detector and HP ChemStation Rev.A.07.01 software for data collection and analysis was used. Separation was carried out on a 250 mm × 4.6 mm i.d., 5 ␮m, reversed-phase Luna C18 column (Phenomenex, Torrance, CA, USA). Solvent A was deionized water containing 0.1% formic acid, and solvent B was acetonitrile with 0.1% formic acid. A linear solvent gradient was used, changing from 30% B to 70% B in 30 min, then to 100% B in 3 min. After 6 min of eluting the column with 100% B, the initial 30% B were reached in 1 min and kept for 4 min before the next injection. The flow rate was 1 ml/min. Preparative HPLC was carried out on a Shimadzu CBM-20A with a 250 mm × 8 mm i.d, 5 ␮m, reversed-phase Knauer Vertex Eurosphere 100-C18 column. Solvent A was deionized water with 0.1% formic acid and solvent B was acetonitrile with 0.1% formic acid. With a flow rate of 4 ml/min, the linear solvent gradient started at 30% B, changed to 70% B in 25 min and then to 100% B in 2 min. After another 2 min at 100% B, the gradient returned to the initial 30% B within 1 min. The detector was set to 270 nm. Under these conditions, the Rt was as follows: ␤-ZAL 13.5 min, ␣-ZAL 16.0 min, ZAN 19.5 min and ZAN-7-dioxolan 21.7 min. HPLC fractions were concentrated under reduced pressure to remove the acetonitrile, and the separated compounds were extracted from the aqueous solutions with t-BME. 2.6. LC–DAD–MS analysis A LXQ Linear Ion Trap MSn system (Thermo Fisher Scientific Inc., Waltham, MA, USA) equipped with a Finnigan Surveyor Autosampler Plus and a Finnigan Surveyor PDA Plus detector was used. This allowed on-line detection of UV absorption and mass spectrometry. The column and solvents were the same as with HPLC. A linear solvent gradient was used, changing from 30% B to 100% B in 30 min. After 1 min of eluting the column with 100% B, the initial 30% B were reached in 1 min and kept for 4 min before the next injection. The flow rate was 0.5 ml/min. The mass spectrometer was operated in the negative electrospray ionization (ESI) mode. Nitrogen was used as sheath gas and auxiliary gas with flow rates of 30.0 and 15.0 l/min, respectively. Spray voltage was 4.5 kV and capillary temperature was 350 ◦ C. Ion optics were automatically tuned with a 10 ␮M solution of ␣-ZAL in methanol. MSn were conducted at CID 35 (35% of 5 V).

3. Results 3.1. Oxidative metabolites of ˛-ZAL and ZAN Human hepatic microsomes were incubated with ␣-ZAL in the presence of a NADPH-regenerating system and the incubation was subsequently extracted with ethyl acetate. Control incubations without NADPH had shown that more than 95% of ␣-ZAL was recovered under these conditions. The reversed-phase LC–DAD–MS profile of the extract from the complete incubation is depicted in Fig. 4 (left chart). The formation of at least eight products was observed which were not generated in the control incubations without NADPH. The least polar peak had a retention time, ESI-mass spectrum and ESIdaughter ion spectrum (ESI-MS2 ) of the M−H ion identical with that of authentic ZAN (Table 1). The ESI-mass spectra of peaks 1, 2 and 4 exhibited M−H ions at m/z 337, suggesting the structure of monohydroxylated ␣-ZALs, whereas peaks 3, 5 and 8 had M−H ions

at m/z 335, compatible with the structures of monohydroxylated ZANs (Table 1). The ion chromatogram of m/z 335 revealed another metabolite, i.e. peak 7, which eluted together with ␣-ZAL (Fig. 4). Peak 6 had an M−H ion at m/z 333. A very similar pattern of metabolites as obtained from ␣ZAL was observed when ZAN was incubated with human liver microsomes (Fig. 4, right chart). This is due to the fact that ZAN, and possibly its hydroxylated metabolites, are reduced to the respective ␣-ZALs by microsomal hydroxysteroid dehydrogenase (Malekinejad et al., 2005). As the position of the newly introduced hydroxyl group could not be deduced from the mass spectra of the ␣-ZAL metabolites, two deuterated analogs of ␣-ZAL, i.e. 6,6,8,8,13,15-D6 -␣-ZAL and 13,15-D2 -␣-ZAL (Fig. 2), and the respective deuterated analogs of ZAN were subjected to microsomal metabolism. The same pattern of metabolites was obtained as with unlabeled ␣-ZAL and ZAN. However, the M−H ions of four of the monohydroxylation products, i.e. peaks 2, 4, 7 and 8, exhibited the loss of one deuterium when either type of deuterated compound was used, clearly indicating that these metabolites had undergone hydroxylation at the aromatic ring (Table 1). In contrast, metabolites 1 and 5 lost one deuterium when the hexadeuterated compounds were metabolized, but did not lose deuterium from the dideuterated compounds. This can only be explained by assuming that metabolites 1 and 5 are hydroxylated at position 6 or 8. The remaining metabolites 3 and 6 lost two deuterium atoms when generated from hexa- or dideuterated ␣-ZAL. This loss and the M−H ion of metabolite 3 at m/z 335 can best be explained by assuming the structure of a quinone of 13- or 15-hydroxy-␣-ZAL, which would lose one deuterium during hydroxylation and the second deuterium through keto-enol tautomerization. Likewise, metabolite 6 may have the structure of a quinone of 13- or 15-hydroxyZAN. Although the use of the deuterium-labeled compounds provided compelling evidence that the preferred site for the hydroxylation of ␣-ZAL and ZAN by human hepatic microsomes was position 13 or 15, the data did not allow to differentiate between these sites. Therefore, the 13-hydroxy- and 15-hydroxy-derivatives of ␣ZAL and ZAN were chemically synthesized according to the route depicted in Fig. 3. The starting materials, i.e. 13-nitro-ZEN and 15nitro-ZEN, have recently been synthesized and fully characterized by NMR spectroscopy in our lab (Pfeiffer et al., 2009). Reduction of the pure nitro-ZEN isomers to the respective amino-ZANs, followed by air oxidation to their quinone imines, hydrolysis to quinones, and reduction of the quinones yielded authentic 13-hydroxyZAN and 15-hydroxy-ZAN (Fig. 3). Whereas 13-hydroxy-ZAN was obtained in good yield, the yield of 15-hydroxy-ZAN was less than 10%, possibly due to a higher reactivity of the intermediate quinone imine and/or quinone.

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Fig. 4. LC–DAD–MS profiles of the extracts from the incubations of ␣-ZAL (left) and ZAN (right) with NADPH-fortified human liver microsomes. Top: UV absorbance; below: ion chromatogram of m/z 337 (M−H of hydroxy-␣-ZAL), m/z 335 (M−H of hydroxy-ZAN and hydroxy-␣-ZAL-quinone) and m/z 333 (M−H of hydroxy-ZAN-quinone).

Synthetic 15-hydroxy-ZAN and the ZAN metabolite peak 7 had identical retention time as well as identical ESI-mass spectra and daughter ion spectra (Table 1). Likewise, synthetic 13-hydroxy-ZAN was identical with the ZAN metabolite peak 8 in LC–MS. From the synthetic 13-hydroxy- and 15-hydroxy-ZAN, the respective hydroxylated derivatives of ␣-ZAL and ␤-ZAL were easily accessible by chemical reduction with sodium borohydride. By comparison of retention time, ESI-MS and ESI-MS2 , metabolite 2 was identical with 15-hydroxy-␣-ZAL and metabolite 4 with 13hydroxy-␣-ZAL. During our experiments with the synthetic 13- and 15-hydroxy derivatives of ␣-ZAL and ZAN, it was noted that these compounds are chemically unstable and prone to autoxidation to their respective quinones. The quinone obtained from 13-hydroxy-␣-ZAL had the same retention time, ESI-MS and ESI-MS2 as metabolite 3 in Fig. 4. By the same criteria, metabolite 6 was identical with 13hydroxy-ZAN-quinone. Moreover, it was noted that the catechol

metabolites hydroxylated at C-15 autoxidized faster than those hydroxylated at C-13. Thus, the ratio of 15- to 13-hydroxy metabolites decreased with the “aging” of the samples. In summary, the major oxidative metabolites generated by human hepatic microsomes from ␣-ZAL and its reductive metabolite ZAN are catechols arising from aromatic hydroxylation at positions 13 and 15. These catechols are unstable, as indicated by the detection of two quinone metabolites among the microsomal products. 3.2. Activity of human CYP isoforms for the hydroxylation of ˛-ZAL In order to clarify which isoforms of hCYP are responsible for the formation of the oxidative metabolites of ␣-ZAL depicted in Fig. 4, 10 major isoforms, i.e. 1A1, 1A2, 1B1, 2A6, 2B6, 2C8, 2C9, 2C19, 2D6 and 3A4, were incubated with ␣-ZAL and the metabo-

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Fig. 5. Activities of major human CYP isoforms for the hydroxylation of ␣-ZAL. Data are the mean ± S.D. of three independent experiments.

lites analyzed by HPLC. Metabolite formation was most pronounced with hCYP1A2 and clearly detectable with hCYP3A4, 2C19, 2D6 and 1B1. LC–DAD–MS analysis of the oxidative metabolites of ␣-ZAL generated by hCYP1A2, hCYP1B1, hCYP2D6 and hCYP2C19 showed the exclusive formation of aromatic hydroxylation products. The major metabolites of these isoforms were 13- and 15-hydroxy-␣ZAL, detected in about the same quantity, but a small amount of 13-hydroxy-␣-ZAL-quinone was also formed. hCYP3A4 appeared to be less specific and lead to both aromatic and aliphatic hydroxylation. Interestingly, the only catechol metabolite generated by hCYP3A4 in significant amount was 13-hydroxy-␣-ZAL, with only traces of 15-hydroxy-␣-ZAL detectable even in fresh samples. As the amounts of metabolites was determined under conditions of linear product formation, the activities of the individual hCYP isoforms could be estimated from the sum of all oxidative metabolites. As depicted in Fig. 5, hCYP1A2 had by far the highest activity, i.e. 51.7 ± 3.1 pmol min−1 pmol CYP−1 , whereas the activity of hCYP3A4 was about 10-fold lower. Very low activities were observed for hCYP2C19, hCYP2D6, and hCYP1B1. The activities of the remaining five isoforms were below the detection limit of 0.1 pmol min−1 pmol CYP−1 . The activity of hCYP1A2 was also determined for ZAN and found to be 30.9 ± 6.2 pmol min−1 pmol CYP−1 . 3.3. Reactivity of the catechol metabolites of ˛-ZAL and ZAN with thiols The autoxidation of 13-hydroxy-␣-ZAL and 15-hydroxy-␣-ZAL to their quinones prompted us to probe the reactivity of these

Fig. 6. LC–MS profiles of the incubations of ␣-ZAL with NADPH-fortified human liver microsomes in the presence of NAC. From top to bottom: MS4 of m/z 498 (M−H of the NAC adduct of hydroxy-␣-ZAL); MS4 of m/z 496 (M−H of the NAC adduct of hydroxy-ZAN and hydroxy-␣-ZAL-quinone); MS4 of m/z 494 (M−H of the NAC adduct of hydroxy-ZAN-quinone).

quinones with N-acetylcysteine (NAC) as a model thiol. NAC was added to the microsomal incubation of ␣-ZAL and the incubation medium subsequently analyzed by LC–DAD–MS. A total of six NAC adducts was detected (Fig. 6). Their identification as listed in Table 2 was based on their LC retention times and on their ESI-MS4 by comparison with reference adducts obtained by incubation of the synthetic 13-hydroxy- and 15-hydroxy-derivatives of ␣-ZAL and ZAN with NAC in aqueous buffer. The MS of each adduct was dominated by the M−H ion, together with small intensities of ions exhibiting a sequential loss of 129 amu (most likely CH2 C(COOH)NHCOCH3 ) and 34 amu (H2 S) from the

Table 2 Characterization of the NAC adducts detected by LC–DAD–MS analysis in incubations containing NADPH-fortified human hepatic microsomes, ␣-ZAL and NAC. Peaka

Rt LC–MS (min)

ESIb -MS (M−H)

A B C D

10.1 12.0 12.7 13.4

498 498 496 496

498 < 369 < 335 498 < 369 < 335 496 < 367 < 333 496 < 367 < 333

E

14.9

496

496 < 367 < 333

16.4

494

494 < 365 < 331

F a b c

Designation as in Fig. 6. Negative mode. Collision-induced dissociation.

Sequential CIDc of ions with m/z

ESI-MS4 of M-(N-acetylcysteinyl) (relative intensity)

N-acetylcysteinyl derivative of

335 (3), 317 (7), 307 (100), 291 (12) 335 (3), 317 (7), 307 (100), 291 (52), 263 (8) 333 (11), 315 (18), 305 (100), 289 (19), 261 (5), 191 (9) 333 (33), 315 (17), 305 (39), 289 (100), 265 (13), 261 (6), 245 (12), 217 (8), 191 (7) 333 (3), 315 (16), 305 (100), 289 (59), 271 (7), 261 (6), 191 (3), 124 (5) 313 (18), 303 (100), 287 (64), 269 (7), 259 (5)

15-HO-␣-ZAL 13-HO-␣-ZAL 15-HO-ZAN 13-HO-␣-ZAL-quinone 13-HO-ZAN 13-HO-ZAN-quinone

A. Hildebrand et al. / Toxicology Letters 192 (2010) 379–386

Fig. 7. Proposed structures of the NAC adducts formed from the catechol metabolite 13-hydroxy-␣-ZAL. The chemical structure of the adduct and its quinone is tentative because isomeric structures are possible.

M−H ion). The MS2 , obtained through collision-induced fragmentation of the M−H ion, exhibited increased intensities of these fragment ions (data not shown). When the fragment ion containing the ZAL or ZAN part of the adduct without the sulfur, obtained by MS3 , was subjected to further collision-induced fragmentation, a fragment pattern was obtained which was specific for the exact structure of the ZAL-derived moiety. Thus, adducts hydrox-

385

ylated at C-13 could be distinguished from their C-15 isomers (Table 2). As shown in Fig. 6 and Table 2, the microsomal incubation of ␣-ZAL with human hepatic microsomes in the presence of NAC gave rise to the NAC adducts arising from the catechol metabolites of ␣-ZAL and its microsomal metabolite ZAN after oxidation to their respective quinones. Quinone formation may have occurred through autoxidation or CYP-mediated oxidation of the catechols. The tentative structure of the adduct derived from 13-hydroxy-␣ZAL is depicted in Fig. 7 (left formula). Small amounts of the primary adducts formed from the quinones and NAC were further oxidized to their quinones (Fig. 7, right formula), as two of such adducts were detected in the incubations with NAC (Table 2 and Fig. 6). When ␣-ZAL was incubated with hCYP1A2 in the presence of NAC, the adducts arising from both catechol metabolites of ␣-ZAL were detected by LC–DAD–MS, together with small amounts of 13hydroxy-␣-ZAL-quinone (data not shown). Only trace amounts of the respective ZAN adducts were detectable, probably due to the fact that supersomes lack hydroxysteroid dehydrogenase and are therefore unable to generate ZAN from ␣-ZAL. In summary, the formation of several NAC adducts derived from the catechol metabolites of ␣-ZAL and its reductive metabolite ZAN clearly indicate that electrophilic intermediates, e.g. quinones, are involved in the catechol pathway. Because different isomeric quinones and possibly quinone methides may be formed from the catechol metabolites, the elucidation of the exact structures of the quinones and their NAC adducts requires further studies. 3.4. Species differences in the microsomal metabolism of ˛-ZAL In order to clarify whether the aromatic hydroxylation of ␣ZAL and its metabolite ZAN is restricted to human CYPs, ␣-ZAL was incubated with rat, bovine and porcine liver microsomes and the microsomal metabolites analyzed by LC–DAD–MS. The profiles obtained are depicted in Fig. 8. As with human hepatic microsomes, ZAN was a major metabolite with each species. The catechol metabolites of ␣-ZAL (metabolites 2 and 4) and ZAN (metabolites 7 and 8) were also detected in all microsomal incubations; however, catechol formation appeared to be less pronounced than in human microsomes. Instead, notable amounts of aliphatic monohydroxylation products of ␣-ZAL and ZAN were observed, the precise chemical structures of which must be elucidated in further studies. 4. Discussion

Fig. 8. Pattern of ␣-ZAL metabolites generated by rat, bovine and porcine liver microsomes. a and b are metabolites hydroxylated at the aliphatic ring.

Although ␣-ZAL is used as a hormonally active growth promotor in cattle for more than two decades, few studies have been conducted on its phase I metabolism. Ingerowski and Stan (1979) reported the oxidation of ␣-ZAL to ZAN in bovine liver, muscle and uterine tissue in vitro, and the same metabolite was demonstrated together with small amounts of ␤-ZAL in microsomes and cytosol preparations from lamb liver by Pompa et al. (1988). ZAN was also observed as a major metabolite of ␣-ZAL in vivo in the urine of female Wistar rats, New Zealand rabbit, beagle dog, rhesus monkey and man, together with small amounts of ␤-ZAL in rabbit urine (Migdalof et al., 1983). Because tritium-labeled ␣-ZAL was used in the in vivo metabolic study, another more polar phase I metabolite could be detected in human urine and was tentatively identified as monohydroxylated ␣-ZAL by gas chromatography–mass spectrometry (GC–MS) after trimethylsilylation. However, the exact structure of this metabolite was not elucidated. Kim et al. (1986) reported ␤-ZAL and small amounts of ZAN as urinary metabolites of bulls dosed with ␣-ZAL. More recently, the phase I metabolites of ␣-ZAL generated by rat and bovine hepatic microsomes were analyzed by HPLC and GC–MS in our laboratory (Metzler and Pfeiffer, 2001). In addition to ZAN and ␤-ZAL, up to five monohydroxy-

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lated metabolites of ␣-ZAL were tentatively identified by GC–MS analysis, but the exact chemical structures could not be clarified. Our interest in the phase I metabolism of ␣-ZAL was rekindled by the recent observation, in our laboratory, that aromatic hydroxylation is a major metabolic pathway of the mycotoxin ZEN with human hepatic microsomes (Pfeiffer et al., 2009). The data presented in this communication provide compelling evidence that aromatic hydroxylation also constitutes a prominent phase I biotransformation of the growth promotor ␣-ZAL and its major metabolite ZAN. The only clue to catechol formation of ␣ZAL and ZAN prior to this report has been published by Li et al. (1985) in the proceedings of a conference. A radioenzymatic assay was used, in which catechols generated by microsomal hydroxylation are trapped as radioactive metabolites by O-methylation with catechol-O-methyltransferase, using the cosubstrate S-adenosyll-methionine with a radiolabeled methyl group as the methyl donor. When ␣-ZAL and ZAN were incubated with liver and kidney microsomes of Syrian hamsters under these conditions, a single radioactive peak was observed with each compound in HPLC analysis, thought to be the monomethyl ether of a catechol metabolite. The radioactive catechol monomethyl ether of ZAN from liver microsomal incubations was further analyzed by gas chromatography–mass spectrometry after trimethylsilylation, and the molecular ion observed in the electron impact mass spectrum was compatible with the structure of a methoxy-ZAN (Li et al., 1985). With the disclosure of catechol formation as a major metabolic pathway of ␣-ZAL and ZAN, the biotransformation of these highly estrogenic compounds becomes more similar to that of the steroidal estrogens 17␤-estradiol (E2) and estrone (E1), the molecules of which also consist of one aromatic ring and an extended aliphatic region. E2 and E1 form two catechols each, which are further metabolized to monomethyl ethers by the enzyme COMT and by conjugation with sulfate and glucuronic acid (Zhu and Conney, 1998). For the catechols of ␣-ZAL and ZAN, preliminary studies using COMT and LC–MS analysis have also shown the formation of monomethyl ethers, but their exact chemical structures remain to be elucidated (Hildebrand, unpublished data). Further similarities but also notable differences between the catechol formation of the xenoestrogen ␣-ZAL and the endogenous steroid estrogens E2 and E1 are reflected by the human CYP isoforms catalyzing this metabolic pathway. For ␣-ZAL, hCYP1A2 is by far the most active isoform, whereas the activity of hCYP3A4 is one order of magnitude and the activities of hCYP1B1, 2C19 and 2D6 are two magnitudes lower (Fig. 5). For E2 and E1, hCYP1A2 is also the most active isoform, but the activity of hCYP1A1 is only by a factor of 2–5 lower, and hCYP1B1, 3A4, 2C9 and others contribute significantly to the hydroxylation of E2 and E1 (Lee et al., 2003). For ␣-ZAL, the dominating activity of hCYP1A2, which is highly expressed in the liver, and the lack of activity of the extrahepatic hCYP1A1 imply that catechol formation of ␣-ZAL occurs predominantly in the liver rather than in extrahepatic tissues. The similarities and differences in the metabolism of ␣-ZAL and E2, especially in estrogen target organs, should be of particular interest in future studies. Another interesting aspect of the catechol metabolites of ␣-ZAL and ZAN relates to their potential toxicological properties. There is increasing evidence that the catechol metabolites of E2 and E1 contribute to the established carcinogenicity of these steroid estrogens for the female breast and uterus, and it is believed that the catechol metabolites induce genotoxic effects such as oxidative DNA damage and covalent DNA adducts in target tissues after oxidation to the respective quinones (Cavalieri and Rogan, 2006; Bolton and Thatcher, 2008). Although there is no evidence to date that ␣-ZAL is carcinogenic, our study has shown that the catechol metabolites of ␣-ZAL and ZAN are unstable and readily oxidized to reactive quinones capable of forming covalent adducts with N-

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