Bioactivation of Tamoxifen by Recombinant Human Cytochrome P450

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Bioactivation of Tamoxifen by Recombinant Human Cytochrome P450 Enzymes Lisa M. Notley, Cornelia J. F. de Wolf, Rebecca M. Wunsch, Roy G. Lancaster, and Elizabeth M. J. Gillam* Department of Physiology and Pharmacology, School of Biomedical Sciences, University of Queensland, St.Lucia, Australia 4072 Received February 26, 2001

Tamoxifen is a major drug used for adjuvant chemotherapy of breast cancer; however, its use has been associated with a small but significant increase in risk of endometrial cancer. In rats, tamoxifen is a hepatocarcinogen, and DNA adducts have been observed in both rat and human tissues. Tamoxifen has been shown previously to be metabolized to reactive products that have the potential to form protein and DNA adducts. Previous studies have suggested a role for P450 3A4 in protein adduct formation in human liver microsomes, via a catechol intermediate; however, no clear correlation was seen between P450 3A4 content of human liver microsomes and adduct formation. In the present study, we investigated the P450 forms responsible for covalent drug-protein adduct formation and the possibility that covalent adduct formation might occur via alternative pathways to catechol formation. Recombinant P450 3A4 catalyzed adduct formation, and this correlated with the level of uncoupling in the P450 incubation, consistent with a role of reactive oxygen species in potentiating adduct formation after enzymatic formation of the catechol metabolite. Whereas P450s 1A1, 2D6, and 3A5 generated catechol metabolite, no covalent adduct formation was observed with these forms. By contrast, P450 2B6, 2C19, and rat liver microsomes catalyzed drug-protein adduct formation but not catechol formation. Drug protein adducts formed specifically with P450 3A4 in incubations using membranes isolated from bacteria expressing P450 3A4 and reductase, as well as in reconstitutions of purified 3A4, suggesting that the electrophilic species reacted preferentially with the P450 enzymes concerned.

Introduction Tamoxifen is a synthetic antiestrogen used successfully for many years in the treatment of breast cancer. Recently, trials have been conducted to examine the efficacy of tamoxifen treatment in the prevention of breast cancer in women with a strong family history of the disease (1). However tamoxifen treatment is associated with a slight but statistically significant elevation in risk of endometrial cancer which complicates the use of tamoxifen in disease-free women (2). Tamoxifen appears to be a more effective carcinogen in rats, inducing liver cancers (reviewed in ref 3). Since many chemicals exert carcinogenic effects by being metabolized to chemically reactive derivatives, it has been proposed that tamoxifen may act as a carcinogen via its metabolism to genotoxic derivatives. Tamoxifen is known to be metabolized by cytochrome P450 enzymes to several primary metabolites, principally 4-hydroxytamoxifen, N-desmethyltamoxifen, R-hydroxytamoxifen, and 4′-hydroxytamoxifen (4-6). In addition, flavin-containing monooxygenases produce tamoxifen N-oxide (7). Several secondary phase I metabolites may also be generated as a result of the sequential reactions catalyzed by P450s with or without the participation of flavin-containing monooxygenases (4-hydroxytamoxifenN-oxide, N-desmethyltamoxifen-N-oxide and R-hydroxy* To whom correspondence should be addressed. Phone: 61-7-3365 1410. Fax: 61-7-3365 1766. E-mail: [email protected].

tamoxifen-N-oxide, N,N-didesmethyltamoxifen, 4-hydroxyN-desmethyltamoxifen, R-hydroxy-N-desmethyltamoxifen 3,4-dihydroxytamoxifen (tamoxifen catechol), and R,4dihydroxytamoxifen) (8, 9). Tamoxifen is believed to be bioactivated by several enzyme systems to reactive species capable of damaging DNA. DNA damage has been detected by postlabeling analysis of DNA extracted from livers of rats treated with tamoxifen (10) and more recently from the endometria of women who have undergone tamoxifen therapy (11, 12). Various studies have suggested the involvement of Phase II metabolism, particularly sulfonation following initial R-hydroxylation (13, 14), in the carcinogenic process. However other, mechanisms of bioactivation of tamoxifen in human tissues remain to be fully examined. Tamoxifen undergoes metabolism-dependent covalent adduct formation with protein during in vitro incubation with human or rodent liver microsomes. Protein adduct formation has been proposed as a surrogate end-point for the production of reactive species capable of binding to DNA; however, different electrophilic products of tamoxifen bioactivation may be associated to greater or lesser extents with protein and DNA adducts, depending on their intrinsic reactivity or stability and the accessibility of the target macromolecules. Kupfer and colleagues have proposed that drug-protein covalent adduct formation in liver microsomes may follow principally from catechol formation (15) (Scheme 1) and have shown a role for P450s 3A4 and 2D6 in catechol production and, by

10.1021/tx0100439 CCC: $22.00 © 2002 American Chemical Society Published on Web 04/17/2002

Tamoxifen Bioactivation by Cytochrome P450

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Scheme 1. Tamoxifen Bioactivation Pathways Discussed in the Present Studya

a

The position of the tritium radiolabel is shown by an asterisk (*).

extension covalent adduct formation (16). However, the catalysis of drug-protein adduct formation by recombinant P450s was not directly assessed. The principal objectives of the current study were to examine the relative contribution of a range of cytochrome P450 enzymes to the generation of protein adducts in humans, and to characterize the protein targets of covalent binding.

Experimental Procedures Materials. Tamoxifen has been identified as a human endometrial carcinogen following long term use and should be handled with care. N-methyl-[3H]tamoxifen was obtained from NEN Life Science Products (Boston, MA) at a specific radioactivity of 85.5 Ci/ mmol. Metabolite standards for HPLC analyses were kindly provided by Dr J. L. Bolton (University of Illinois at Chicago; catechol, metabolite E, R-hydroxytamoxifen), Dr. I. N. H. White (MRC Toxicology Unit, University Of Leicester, U.K.; 4-hydroxytamoxifen, N-desmethyltamoxifen, N,N-didesmethyltamoxifen, tamoxifen N-oxide), Dr. D. H. Phillips (Institute of Cancer Research, Sutton, U.K.; R-hydroxytamoxifen and R,4-dihydroxytamoxifen), and Dr. M. S. Lennard (University of Sheffield, U.K.; 4′-hydroxytamoxifen). N,N-didesmethyltoremifene HCl was the generous gift of Orion Farmos Corporation (Turku, Finland). Tamoxifen citrate was purchased from the Sigma Chemical Company (St. Louis, MO). All other chemicals were obtained from local suppliers at the highest quality commercially available. Liver Microsomes. Samples of human liver were obtained from organ donors according to procedures approved by University of Queensland ethics committees and frozen in liquid nitrogen for storage at -70 °C prior to use. Microsomes were prepared according to the method of Guengerich (17), with the addition of a final wash in 10 mM Tris-acetate, 1 mM EDTA, 20% glycerol to remove residual drugs. Liver microsomes were also prepared from untreated male Wistar rats (150-200 g; University of Queensland Central Animal Breeding Facility). Cytochrome P450 concentrations were determined as previously described (17). Recombinant P450 Preparations. Human P450 enzymes were expressed in bacteria in bicistronic format with NADPHcytochrome P450 reductase (hNPR)1 as described previously (18-21). DH5R strain Escherichia coli were transformed with bicistronic expression constructs each of which contained cDNAs

encoding hNPR and one of the following recombinant P450s: 1A1; 1A2; 1B1 [four variants, designated RAVN (allele CYP1B1*3), RALN (CYP1B1*1), GSVN (CYP1B1*6), GSLN (CYP1B1*2) expressing the following amino acids at positions 48 (Arg or Gly), 119 (Ala or Ser), and 432 (Val or Leu); all four variants contained Asn at position 453 (22)]; 2A6; 2B6; 2C9*1 (wild-type); 2C19; 2D6 [full-length variant designated DB4 (23), but with wild type (Ala) at position 374]; 2E1; 3A4; 3A5; and 3A7. Cells were also transformed with the monocistronic expression vector containing the cDNA for hNPR alone and with the empty vector, pCW. Bacteria were cultured and harvested as described previously (24) except that the expression of P450 forms 1A1, 2A6, 2B6, 2D6, and 3A7 was augmented by coexpression of bacterial chaperones. Briefly, E. coli strain DH5R were cotransformed with the relevant bicistronic expression vector and the chaperone expression vector pGro7 [a generous gift of Professor K. Nishihara, HSP Research Institute, Kyoto, Japan (25)] using the method of Inoue et al. (26). Colonies harboring both plasmids were selected by growth on LB agar containing 100 µg/mL ampicillin and 20 µg/mL chloramphenicol. Isolated colonies were precultured at 37 °C overnight in LB media containing both antibiotics, then used to inoculate terrific broth containing 1 mM thiamine, trace elements, 100 µg/mL ampicillin, and 20 µg/mL chloramphenicol. Cultures were incubated for 5 h at 25 °C, 160-180 rpm shaking speed before initiation of induction by the addition of arabinose (4 mg/mL), IPTG (1 mM), and δ-aminolevulinic acid (0.5 mM). Flasks were then incubated for a further 43 h at 25 °C, 160-180 rpm before harvest. Membranes were prepared and characterized for P450 hemoprotein expression and hNPR activity. Bacterial membranes were used directly in enzyme assays. Purified Enzyme Preparations. Recombinant cytochrome P450 forms were expressed in bacteria and purified as previously described (24, 27). NADPH-cytochrome P450 reductase and cytochrome b5 were purified by slight modifications of published methods (17). Quantitative Assessment of Covalent Adduct Formation. Incubations contained 0.05-0.5 µM P450 from microsomes or bacterial membranes, 100-150 mM Tris HCl, pH 7.4, supplemented with substrates added from ethanolic or methanolic stocks. Final concentrations of solvent were maintained below 1% v/v in all cases; methanol and ethanol were shown to have the least inhibitory effect on the activities measured (results not shown). Reactions were initiated by the addition of 1 Abbreviations: HL, human liver; hNPR, human NADPH-cytochrome P450 reductase; RL, rat liver; ROS, reactive oxygen species.

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an NADPH generating system consisting of (final concentrations) 1 mM NADPH, 2.5 mM glucose-6-phosphate, and 0.5 units/mL glucose-6-phosphate dehydrogenase. Reaction mixtures were incubated at 37 °C with gentle agitation for 60 min unless otherwise indicated. In some incubations, levels of hNPR were supplemented by adding additional membranes from cells expressing hNPR alone to a reductase concentration five times that of the P450 concentration for P450 2C forms or up to two times the level of P450 used for other forms. Incubations were also supplemented with purified cytochrome b5 at an equal molar ratio to the P450 concentration where indicated. Incubations were undertaken under reduced light and using amber glass and plasticware to minimize photodegradation of tamoxifen. In certain cases, catalase (100 µg/mL) or superoxide dismutase (SOD; 15 µg/mL) were added, or the concentration of NADP+ in the regenerating system was reduced to 0.2 mM (and consequently the concentration of NADPH generated was also 0.2 mM). Adduct formation was assayed by liquid scintillation counting of tritium remaining associated with protein after extraction with detergents and solvents as described by Munns et al. (28), to remove unbound radiolabel. Total radiolabel concentrations in incubations were confirmed by scintillation counting of aliquots of the residual reaction mixtures (that were not subjected to extraction). Fluorography. Aliquots from incubations of bacterial membranes or microsomes with radiolabeled tamoxifen were quenched by the addition of 0.6 volumes of gel loading buffer (181 mM Tris HCl, pH 6.8, containing 45% v/v glycerol, 2.7% w/v SDS, 0.0027% w/v bromophenol blue, and 13% v/v 2-mercaptoethanol), and subjected directly to SDS-PAGE according to the general method of Laemmli (29) in 8.3% polyacrylamide gels. Fluorography was carried out as described by Munns et al. (28). Experiments with purified recombinant P450 3A4 were carried out in a similar fashion except that P450 3A4 was reconstituted with purified human reductase and b5 (24). Substrate concentrations and assay durations were as noted in the results for individual experiments. HPLC Analysis of Tamoxifen Metabolism to the Catechol Metabolite. All procedures were undertaken under reduced light and using amber glass and plasticware to minimize photodegradation of tamoxifen. Incubations were performed as above but with 0.05-0.2 µM P450 from microsomes or bacterial membranes, 100 µM trans-4-hydroxytamoxifen, and an NADPH-generating system in 100 mM potassium phosphate, pH 7.4, with 2 mM ascorbic acid unless otherwise noted in the legends of figures showing results of individual experiments. Incubations using P450 3A and 2C forms were supplemented as described above with membranes obtained from cells expressing hNPR alone, unless otherwise noted. Control incubations were included in each assay run, and contained membranes from cells expressing hNPR alone (“hNPR”) or from cells transformed with the empty expression vector (“pCW”). Incubations were allowed to proceed for 120 min before being terminated by addition of 2 mL of helium-purged ethyl acetate, followed by addition of 100 µL of 15 µM didesmethyl toremifene HCl (internal standard). The phases were mixed vigorously then the top 1.4 mL were removed. A second extraction was then performed with a further 1.2 mL of ethyl acetate. The combined extracts were evaporated under argon then resuspended in 100 µL of helium-purged acetonitrile prior to analysis by HPLC. HPLC was performed using a Shimadzu HPLC system fitted with an auto-injector and a 3.9 × 150 mm Waters Symmetry C8 reverse phase column. The mobile phase was 20 mM ammonium acetate: acetonitrile run on a gradient derived from that described in Poon et al. (9), and modified by extension of the second gradient step to optimize peak separation. Initial conditions were 95:5 20 mM ammonium acetate:acetonitrile. The following steps were programmed: 0-4 min linear gradient to 80:20 ammonium acetate:acetonitrile; 4-24 min linear gradient to 60:40 ammonium acetate:acetonitrile; 24-60 min linear gradient to 35:65 ammonium acetate:acetonitrile; 60-70 min constant at 35:65 ammonium acetate:acetonitrile; 70-80 min linear gradient to 95:5 ammonium acetate:acetonitrile; and a

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Figure 1. Covalent drug-protein adduct formation from tamoxifen assessed using a range of recombinant human P450 preparations. Bacterial membranes containing recombinant P450s coexpressed with hNPR or microsomes from untreated male rat liver (RL) or human liver (HL) were incubated at a P450 concentration of 0.15 µM with 28.2 nM [3H]tamoxifen. Incubations containing P450 3A and 2C forms were tested both with and without supplementation of membranes obtained from cells expressing hNPR alone. Control incubations contained membranes from cells expressing hNPR alone (“hNPR”), and from cells transformed with the empty expression vector (“pCW”). Reactions were quenched after 60 min and radiolabel covalently associated with protein was quantified as described in Experimental Procedures. Solid bars represent complete incubations and empty bars show NADPH-deficient controls. Results are the mean ( SD of triplicate determinations. Asterisks denote results significantly different to NADPHdeficient controls (Student’s t test): (*) p < 0.05; (**) p < 0.01; (***) p < 0.001. final reequilibration at 95:5 ammonium acetate:acetonitrile for 10 min. The flow rate was 0.75 mL/min. Metabolites were detected by absorbance at 280 nm. Under these conditions the retention times of tamoxifen and its metabolites were as follows: R,4-dihydroxytamoxifen, 28.4 min; R-hydroxytamoxifen, 36.3 min; 3,4-dihydroxytamoxifen (catechol) isomers 38.6/39.0 min; trans-4-hydroxytamoxifen, 44.4 min; cis-4-hydroxytamoxifen, 46.1 min; 4′-hydroxytamoxifen, 46.9 min; N,N-didemethyltamoxifen, 53.1 min; N-desmethyltamoxifen, 56.1 min; metabolite E isomers, 61.4/64.2 min; tamoxifen N-oxide, 62.4 min; tamoxifen, 62.3 min. Other Methods. Hydrogen peroxide production was measured as described by Bell and Guengerich (30). Midazolam hydroxylation and nifedipine oxidation were measured by the general methods described previously (17, 31). Activity was measured using 100 µM midazolam or 200 µM nifedipine, and 0.1-0.25 µM P450 for 5 min, conditions under which product formation increased linearly with respect to time and enzyme concentration.

Results 3

Incubation of [ H]tamoxifen with bacterial membranes containing coexpressed cytochrome P450 forms and NADPH-cytochrome P450 reductase revealed significant cofactor-dependent covalent binding of radiolabel to protein catalyzed by recombinant P450s 3A4 and, to a lesser extent, P450s 2B6 and 2C19 (Figure 1). Covalent adduct formation was also seen in rat liver microsomes and at low levels in a range of human liver microsomes; however, levels of activity were considerably lower than anticipated from the activity of recombinant P450 3A4 (Figure 2). Moreover adduct formation did not appear to correlate with P450 3A activity as measured by nifedipine oxidation or midazolam 1′- or 4-hydroxylation (r2 ) 0.008,


Figure 2. Examination of covalent drug-protein adduct formation from tamoxifen in human liver microsomes. Incubations were conducted with human and rat liver microsomes and recombinant P450 3A4/hNPR preparations (not supplemented with additional reductase or cytochrome b5) as described in the Experimental Procedures using 0.15 µM P450 and 24 nM [3H]tamoxifen. Reactions were quenched after 60 min, and radiolabel covalently associated with protein was quantified as described in Experimental Procedures. Numbers denote individual human liver preparations. RL signifies liver microsomes derived from a single untreated male rat. Four separate individual male rat liver preparations incubated under identical conditions showed a mean (( SD) of 80 ( 48 fmol tamoxifen equivalents bound/ml incubation corrected for radiolabel association seen in NADPH-deficient controls. Incubations labeled 3A4 and pCW denote incubations containing membranes from bacteria transformed with the bicistronic expression vector for P450 3A4 and hNPR and with the empty pCW vector, respectively. Solid bars represent complete incubations and crosshatched bars show NADPH-deficient controls. Results are the mean ( SD of triplicate determinations. Asterisks denote results significantly different to NADPH-deficient controls (Student’s t test) at the following levels: (*) p < 0.05; (**) p < 0.01; (***) p < 0.0001.

0.001, and 0.022, respectively), yet strong correlations were seen between these classical markers of P450 3A activity (midazolam 1′- vs 4-hydroxylation, r2 ) 0.838; midazolam 1′-hydroxylation vs nifedipine oxidation, r2 ) 0.926; and midazolam 4-hydroxylation vs nifedipine oxidation, r2 ) 0.895, respectively, over eight liver preparations). Addition of native or heat-treated liver microsomes failed to decrease P450 3A4-mediated adduct formation to any greater extent than comparable amounts of bacterial membranes (Figure 3). The addition of cytochrome b5 reduced adduct formation catalyzed by P450 3A4 to a limited extent; however, supplementation of P450 3A4/hNPR membranes with additional reductase decreased adduct formation markedly (Figure 4). Adduct formation in a range of preparations was seen to correlate inversely with the hNPR:P450 3A4 ratio (Figure 5A) but not overall protein concentration (Figure 5B). Recombinant P450 3A4 preparations showed significantly elevated hydrogen peroxide production that could be reduced to baseline levels by reductase supplementation (Figure 6A). By comparison, hydrogen peroxide was markedly lower in a selection of human liver microsomes incubated under identical conditions (Figure 6B). Tamoxifen 3,4-catechol was formed from 4-hydroxytamoxifen in recombinant preparations of P450 3A4 and to a lesser extent P450s 3A5, 2D6, and 1A1 (Table 1) as well as with human liver microsomes (Table 2). No catechol could be detected in incubations with other recombinant forms or rat liver microsomes (data not shown). Two peaks could be resolved in extracts derived from incubations with P450 3A4 that were tentatively designated as the cis and trans isomers of the catechol.


Figure 3. Effect of addition of human liver microsomes or bacterial membranes on P450 3A4-dependent covalent binding of radiolabel derived from tamoxifen. Incubations were conducted with recombinant P450 3A4/hNPR (not supplemented with additional reductase or cytochrome b5) as described in the Experimental procedures using 0.15 µM P450 and 24 nM [3H]tamoxifen for 60 min. 3A4 + HL and 3A4 + pCW signify P450 3A4/hNPR incubations containing human liver microsomes (HL27) and membranes from bacteria transformed with the empty pCW vector, respectively, and total protein concentrations were made up to 0.55 mg/mL in both cases. Controls (HL and pCW) contained only the equivalent amount of HL27 microsomes or pCW bacterial membranes. HT signifies addition of microsomes or membranes that had been previously subjected to a 60 °C, 30 min heat treatment. Solid bars represent complete incubations and crosshatched bars show NADPH-deficient controls. Results are the mean ( SD of triplicate determinations.

Though pure standards of isomers were not available, assignment of the peaks was based on the anticipated preferential formation of trans-catechol from trans-4hydroxytamoxifen. Production and/or recovery of the catechol metabolite was reduced by omission of ascorbic acid or additional reductase from P450 3A4 incubations (Table 2); however, addition of ascorbic acid failed to affect catechol production by human liver microsomes. No catechol formation was apparent in incubations of rat liver microsomes with 100 µM 4-hydroxytamoxifen; however, R-hydroxytamoxifen and 4-hydroxytamoxifen were both produced in comparable incubations with 250 µM tamoxifen (data not shown). The potential for uncoupling in other recombinant P450 preparations was assessed by measuring hydrogen peroxide production in the presence and absence of substrate. No other P450s shown to generate tamoxifen catechol demonstrated significant hydrogen peroxide production above levels observed in the presence of hNPR alone (data not shown). The role of ROS and oxidizing conditions in covalent adduct formation was examined in two further experiments. First, the effect of catalase or superoxide dismutase (SOD) on adduct formation by P450 3A4 was assessed. Neither enzyme significantly affected adduct formation (results not shown; p > 0.05, two tailed Students t-test). Second, covalent adduct formation was assessed at a lower NADPH concentration (Table 3). Whereas adduct formation in the presence of a 1mM NADPH generating system for rat liver microsomes was slightly but not significantly higher than at 0.2 mM (p ) 0.06, two tailed Student’s t-test), it was higher at 0.2 mM NADPH for incubations with both a single human liver and recombinant P450 3A4. Importantly, significant adduct formation was seen in incubations with P450 2D6

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Figure 4. Effect of cytochrome b5 supplementation on drugprotein adduct formation from tamoxifen catalyzed by recombinant human P450 preparations. The effect of cytochrome b5 on covalent adduct formation was assessed in a range of bacterially expressed P450 forms previously demonstrating enhanced activity in the presence of cytochrome b5. Incubations were conducted with 0.15 µM P450 and 28.2 nM [3H]tamoxifen, with and without an equal molar addition of purified rabbit b5, and in some cases, with additional hNPR supplementation to two (P450 3A forms and P450 2A6) or five (P450 2C forms) times the P450 concentration. Control incubations contained membranes from cells expressing hNPR alone (“hNPR”). Solid bars represent complete incubations and empty bars show NADPHdeficient controls. Results are the mean ( SD of triplicate determinations. Asterisks denote results significantly different to NADPH-deficient controls (Student’s t test): (*) p < 0.05; (**) p < 0.01; (***) p < 0.001.

with the 0.2 mM but not the 1 mM NADPH generating system. The proteins subject to covalent modification by tamoxifen were characterized using SDS-PAGE and fluorography. Radiolabel was found to localize with multiple bands the molecular weights of many of which corresponded to those generally expected of P450s (45-60 kDa) in incubations with rat liver microsomes (Figure 7A). In bacterial membranes containing recombinant P450 3A4/hNPR (Figure 7B) and in reconstitutions of purified P450 3A4 (Figure 7C), radiolabel was more clearly associated with a predominant band corresponding to P450 3A4 but some radiolabel also comigrated with the low molecular weight protein front and, to a lesser extent, with other proteins in incubation mixtures.

Discussion Previous studies by Kupfer and colleagues have proposed that drug-protein adduct formation from tamoxifen follows from generation of a catechol intermediate by P450s 3A4 and 2D6 (15, 16). Adduct formation was potentiated by using 4-hydroxytamoxifen as substrate (15); P450 3A-selective chemical inhibitors and antibodies decreased covalent binding of radiolabel from radiolabeled tamoxifen in human and rat liver microsomes (32); and recombinant P450 3A4 and to a lesser extent, P450 2D6, catalyzed catechol formation (16). However, no clear correlation was seen in a previous study (32), and only a weak correlation in another (33), between covalent adduct formation and P450 3A-dependent activities in human liver microsomes, prompting a direct examination of the ability of recombinant P450 preparations to catalyze drug-protein adduct formation.

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Figure 5. Correlation of covalent drug-protein adduct formation with hNPR:P450 ratio (A) and P450 specific content (B) in recombinant membrane preparations. Covalent adduct formation was assessed at a concentration of 24 nM [3H]tamoxifen for 60 min. Data represent the mean ( SD of triplicate determinations. Solid circles show the same recombinant P450 3A4/hNPR preparation before and after supplementation with additional hNPR-containing membranes. Data were fitted using logarithmic (A) and linear (B) models generating the R2 values shown on the figures.

In the present study we were able to demonstrate for the first time that recombinant P450 3A4 was able to catalyze covalent drug-protein adduct formation from tamoxifen. In contrast to the prediction that would be made on the basis of studies of catechol formation (16), P450 2D6 failed to catalyze significant adduct formation under standard assay conditions (1 mM NADPH generating system). As in the study by Mani et al. (32), covalent adduct formation in liver microsomes failed to correlate with three different P450 3A marker activities. Indeed livers shown to have significant P450 3A4dependent activity failed to demonstrate significant covalent adduct formation. Three hypotheses were proposed to explain this lack of correlation. First, we hypothesized that some component present in human liver microsomes inhibited covalent adduct formation. This was tested by adding untreated and heattreated microsomes from a liver showing low activity to incubations with recombinant P450 3A4/hNPR membranes. The effect of bacterial membranes was tested in parallel as a control. Whereas addition of any membrane preparations led to a decrease in activity, no differences were seen between liver microsomes and bacterial membranes or between native and heat-treated preparations (Figure 3), suggesting that there was no endogenous inhibitor of adduct formation in liver microsomes. The decrease in adduct formation caused by addition of either type of lipid membrane can be explained by the fact that


Chem. Res. Toxicol., Vol. 15, No. 5, 2002 619 Table 1. Catechol Production by Recombinant P450 Forms and Liver Microsomes P450 preparationa

catechol formation (arbitrary units of peak area)b

1A1 2D6 3A4 3A5

70 ( 10 100 ( 20 3600 ( 700 1000 ( 200

a Bacterial membranes containing recombinant P450s (0.2 µM P450) coexpressed with reductase were used in incubations. In all cases, incubations containing P450 3A forms were supplemented with membranes obtained from cells expressing hNPR alone to a 2-fold molar excess of hNPR over P450, and with purified b5 at an equal molar concentration. Control incubations contained membranes from cells expressing hNPR alone supplemented with purified b5 and from cells transformed with the empty expression vector. b Incubations were carried out with 100 µM trans-4hydroxytamoxifen in the presence of 2 mM ascorbate for 120 min. No coeluting peaks were seen in t ) 0 min controls. Data represent the mean ( SD of three determinations. All data were significantly different to controls (Student’s t test) at the level of p < 0.001. No other recombinant P450 tested, nor either of the controls, yielded detectable levels of catechol.

Table 2. Effect of HNPR Supplementation and Ascorbic Acid on Catechol Recovery from Incubations with Recombinant P450 3A4 and Human Liver Microsomes catechol formation (arbitrary units of peak area)b incubation conditionsa

Figure 6. Hydrogen peroxide production in incubations with P450 3A4/hNPR and human liver microsomes. (A) Effects of redox partner supplementation on uncoupling of P450 3A4/ hNPR preparations. (B) Comparison of uncoupling in recombinant P450 3A4/hNPR preparations and representative human liver microsomes. Incubations were conducted with recombinant P450 3A4/hNPR preparations or human liver microsomes at a P450 concentration of 0.2 µM, and in the presence and absence of an NADPH-generating system and the prototypical P450 3A4 substrate, midazolam (100 µM). At 30 min, incubations were quenched and hydrogen peroxide production was assessed colorimetrically as described under Experimental Procedures. Results are the mean ( SD of triplicate determinations. Asterisks denote results significantly different to NADPHdeficient, substrate deficient controls (Student’s t test): (*) p < 0.05; (**) p < 0.01; (***) p < 0.001. (A) Incubations were undertaken with and without supplementation of recombinant P450 3A4/hNPR membranes obtained from cells expressing hNPR alone and purified b5 as indicated. (B) Incubations were conducted as in (A) but without hNPR or b5 supplementation. HL numbers denote microsomal preparations from individual human livers.

as a lipophilic substrate, tamoxifen is predominantly distributed within the lipid bilayer environment. Thus, addition of extra membranes increases the volume through which the fixed amount of tamoxifen can distribute, lowering its effective concentration. Since the concentration of tamoxifen used in covalent binding assays is likely to be far less than the Km for this process, a decrease in overall adduct formation is to be expected. Second, we proposed that the presence of cytochrome b5 in liver microsomes shifted metabolism of tamoxifen away from the pathway leading to the reactive intermediate. This was tested by adding purified cytochrome b5 to the recombinant system for those forms shown previously to be affected by the presence of b5 (19, 24, 27, 34, 35). Cytochrome b5 slightly decreased adduct formation by P450 3A4 (Figure 4); however, adduct formation was

P450 3A4 complete system (with both hNPR supplementation and ascorbate) P450 3A4 minus hNPR supplementation P450 3A4 minus ascorbate P450 3A4 minus both hNPR supplementation and ascorbate human liver microsomes plus ascorbate human liver microsomes minus ascorbate

putative isomer 1

putative isomer 2

3350 ( 130

750 ( 2

1810 ( 210*

350 ( 30***

2220 ( 230*

890 ( 110

460 ( 20***

340 ( 8***

650 ( 60

170 ( 30

560 ( 20

240 ( 20

a Bacterial membranes containing recombinant P450 3A4 (0.2 µM) coexpressed with hNPR, and human liver microsomes (0.2 µM P450) were incubated with 100 µM trans-4-hydroxytamoxifen for 120 min. As indicated, incubations were supplemented with hNPR from bacterial membranes to a final concentration of 0.4 µM. Ascorbic acid was added where indicated at a final concentration of 2 mM. b Tamoxifen catechol was identified by coelution with an authentic standard. Two closely eluting peaks were seen in both the standard and incubation extracts. Tentative assignment of putative isomer 1 as trans-catechol and of putative isomer 2 as the cis-catechol was based on the assumption that trans-catechol should predominate in incubations using trans-4-hydroxytamoxifen as substrate. Data represent the mean ( SD of three determinations. Asterisks signify data significantly different to fully supplemented controls (plus hNPR and ascorbate for 3A4; plus ascorbate for human liver microsomes) at the following levels: (*) p < 0.05; (**) p < 0.01; (***) p < 0.005 (Student’s t test).

still significantly greater than in human liver microsomes accounting for the level of P450. By contrast, supplementation of recombinant P450 3A4 incubations with additional reductase led to a considerable reduction in adduct formation. This latter observation was consistent with a third hypothesis, that the reducing environment present in microsomal incubations (containing 1 mM NADPH) failed to support oxidation of the catechol to the ultimate

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Table 3. Role of Oxidizing Conditions and ROS on Adduct Formation in Incubations with Recombinant P450 Forms and Liver Microsomesa radiolabel remaining associated with protein after extraction| (fmol of tamoxifen equivalents/mL reaction) incubation conditions

1 mM NADPH generating system

0.2 mM NADPH generating system

NADPH-deficient

RL microsomes HL27 microsomes P450 3A4 P450 2D6

150 ( 3*** 49 ( 2** 38 ( 4* 47 ( 3

104 ( 30* 57 ( 1***† 63 ( 4***† 79 ( 6*†

34 ( 1 37 ( 3 25 ( 4 49 ( 17

a The effect of NADPH concentration on covalent adduct formation by recombinant P450s and liver microsomes was assessed in incubations conducted with 0.2 µM P450 (except 0.013 µM P450 2D6) and 26.1 nM [3H]tamoxifen for 90 min, with and without an NADPHgenerating system at the final NADPH concentrations indicated. The hNPR:P450 ratio in the recombinant P450 incubations was 2 for P450 3A4 and 20 for P450 2D6. Results are the mean ( SD of triplicate determinations. Asterisks denote results significantly different to NADPH-deficient controls (Student’s t test): (*) p < 0.05; (**) p < 0.01; (***) p < 0.001. Daggers denote adduct formation significantly higher than the 1 mM NADPH generating system incubation (Student’s t test): (†) p < 0.005.

Figure 7. Targets of drug-protein adduct formation in rat liver microsomes (A), P450 3A4/hNPR recombinant membrane preparations (B), and reconstitutions of purified recombinant P450 3A4 (C). Enzyme preparations were prepared and incubated under the conditions indicated with 200 nM [3H]tamoxifen for 90 min, then quenched and subjected to denaturing electrophoresis and fluorography, as described in Experimental Procedures. Liver microsomes prepared from an untreated male rat (A) and membranes isolated from bacteria coexpressing P450 3A4 and hNPR (B) were incubated at a P450 concentration of 0.2 µM. Bacterial preparations were not supplemented with additional hNPR. (C) Purified recombinant P450 3A4 (0.2 µM) was reconstituted with and without reductase (0.4 µM), purified rabbit cytochrome b5 (0.2 µM), substrate, an NADPH generating system and reduced glutathione (3 mM). Reactions denoted “t ) 0” were quenched immediately after initiation. Arrows designate the broad molecular weight range corresponding to P450s (45-60 kDa; (A) and the band corresponding to recombinant P450 3A4 (B, C), respectively.

electrophilic products responsible for adduct formation. We proposed that oxidation of NADPH by P450 3A4 was uncoupled from substrate oxidation to a greater degree in recombinant bacterial membranes than in human liver microsomes, leading to the release of reactive oxygen species (ROS) at the P450 active site that could catalyze oxidation of the catechol. This hypothesis was supported by the observation that recombinant P450 3A4 incubated with NADPH led to time-dependent production of hydrogen peroxide, which could be almost abolished by supplementation of incubations with additional hNPR in the same manner as it dramatically reduced drugprotein adduct formation in the same recombinant system (Figure 6A). By contrast, supplementation with b5 only slightly reduced hydrogen peroxide production in the

presence of substrate. This finding was consistent with its effect on adduct formation. Only low levels of hydrogen peroxide production were observed in three representative human liver microsomal preparations incubated concurrently. Indeed the liver microsomal preparation showing the most significant hydrogen peroxide production (HL25; Figure 6B) also showed the most significant adduct formation (Figure 2). In line with this hypothesis, adduct formation correlated inversely with hNPR:P450 ratio in a range of recombinant P450 3A4 preparations (Figure 5A), whereas no significant correlation was observed with P450 specific content and therefore relative protein concentration (Figure 5B). Assuming that the lipid:protein ratio is relatively constant in bacterial membranes, this suggested to us that the effect of reductase supplementation was not simply due to a dilution of substrate in a larger volume of membranes as seen in Figure 3. The addition of SOD and catalase failed to decrease adduct formation, suggesting that oxidation of the catechol occurred within the P450 3A4 active site and was therefore inaccessible to these enzymatic ROS scavengers. Kupfer et al. have shown that P450 3A4, 2D6, and several other forms (2C19, 2C8, 1A2, 1A1, and 2A6) have the potential to produce the catechol metabolite from tamoxifen (16). In the current study, though P450 3A4 was clearly predominant, P450s 2D6, 3A5, and 1A1 were also seen to catalyze catechol formation from 4-hydroxytamoxifen, however no significant adduct formation was observed with these latter forms. Whereas P450s 1A1 and 3A5 do not appear to catalyze significant 4-hydroxylation of tamoxifen and would therefore not generate enough primary metabolite to produce catechol in the covalent binding assay, P450 2D6 has been shown to be a more efficient catalyst of tamoxifen 4-hydroxylation than P450 3A4 (36, 37; Crewe, H. K., Notley, L. M., Wunsch, R. M., Lennard, M. S., and Gillam, E. M. J.).2 It has also been proposed to be competent at 3-hydroxylation of 4-hydroxytamoxifen to the 3,4-catechol (16). No significant hydrogen peroxide formation was noted with this form however, under the conditions used in these assays, suggesting that catechol formation alone is not sufficient to lead to adduct formation; rather ROS are required to perform the final activation step. We predicted that latent potential for adduct formation from catechol formed in P450 2D6 incubations might be revealed in a less reducing environment. Consistent with this hypothesis, adduct formation was observed when P450 2D6 was incubated with [3H]tamoxifen at a lower NADPH concentration. 2

Unpublished results.


Sridar et al. have recently shown that P450 2B6 can catalyze metabolic activation of tamoxifen to a proteinreactive species and that P450 2B6 becomes modified in the process (Sridar, C., Kent, U. M., Notley, L. M., Gillam, E. M. J., and Hollenberg, P. F.).2 We were also able to demonstrate limited adduct formation with recombinant P450 2B6 at the nanomolar substrate concentrations used in the present study. A low but statistically significant level of adduct formation was also seen with P450 2C19. Previous studies have shown that P450 2B6 is apparently a low affinity catalyst of tamoxifen 4-,4′- and R-hydroxylation (Crewe, H. K., Notley, L. M., Wunsch, R. M., Lennard, M. S., and Gillam, E. M. J.),2 but no activity was seen toward catechol formation from 4-hydroxytamoxifen in the present study (results not shown). P450 2C19 catalyzed both N-demethylation and 4-hydroxylation of tamoxifen at high substrate concentrations, as well as trans-cis isomerization of 4-hydroxytamoxifen (Crewe, H. K., Notley, L. M., Wunsch, R. M., Lennard, M. S., and Gillam, E. M. J.),2 but again no activity was detected toward catechol formation (results not shown). Although we cannot exclude the possibility that P450 2B6 or P450 2C19 may catalyze catechol formation using 3-hydroxytamoxifen as substrate, no unidentified peaks that might correspond to 3-hydroxytamoxifen were detected in chromatograms from incubations of tamoxifen with P450 2B6 or P450 2C19. Thus, it seems unlikely that catechol formation underlies the limited drug protein adduct formation observed here. Similarly, we cannot exclude the possibility that 3′4′catechol formation on the other phenyl ring is responsible for adduct formation in these cases; indeed P450 2B6 has been shown to form 4′-hydroxytamoxifen. However, the absence of significant uncoupling with either P450 2B6 or P450 2C19 would also argue against involvement of a catechol metabolite formed on either ring by extrapolation of the observation made with P450 2D6 that catechol formation in the absence of ROS production fails to lead to adduct formation. Covalent adduct formation has been demonstrated previously using rat liver microsomes, and similarly attributed to P450 3A-dependent activity. Whereas the form catalyzing tamoxifen bioactivation in the untreated male liver preparations used here was not identified, it is unlikely that adduct formation proceeds via the catechol metabolite, since no catechol formation was noted using 4-hydroxytamoxifen as substrate (data not shown). We cannot, however, dismiss the possibility that catechol formation proceeds through initial 3-hydroxylation of tamoxifen. By contrast, rat preparations generated both R-hydroxytamoxifen and R,4-dihydroxytamoxifen, both proposed as precursors to DNA adduct formation. Though sulfonation is apparently required to generate a metabolite that is able to form adducts at high levels in vitro (13, 38, 39), R-hydroxytamoxifen is reactive in its own right, albeit to a considerably lesser extent (40). Thus, the protein adduct formation found here with rat liver microsomes may proceed through the R-hydroxylated metabolite. Alternatively, the R,4-dihydroxy metabolite may lead to formation of the quinone methide or tautomers thereof (41), that could covalently bind to protein. Fluorographic analysis of protein adducts formed in incubations with recombinant P450 3A4 in bacterial membranes and reconstitutions of purified P450 3A4 showed selective modification of the P450 band consistent with the P450 catalyst being the chief target of drugprotein adduct formation. In incubations with rat liver


microsomes, bands migrating at molecular weights characteristic of cytochrome P450 enzymes were preferentially labeled, as seen previously in phenobarbital-treated rat liver microsomes (42), though other proteins were also modified to a lesser degree. Thus, in both human recombinant and rat preparations, adduct formation may lead to a preferential modification of the catalytic P450 as seen with many other P450 substrates. Though this effect is unlikely to lead to modification of P450 3A4 under physiological conditions, due to the requirement for oxidizing conditions to potentiate adduct formation, tamoxifen may modify one or more rat P450 forms in the absence of oxidizing conditions. The low level of covalent adduct formation observed with human liver microsomes is likely to reflect the combined contributions of P450s 2B6, 2C19, and P450 3A4 dependent upon the relative expression of the different forms in individual livers and the degree of ROS generation that accompanies P450 3A4 activity. It is notable that a previous study showed a weak correlation of drug-protein adduct formation with P450 2B6 levels in microsomes as well as P450 3A4 levels (33). The requirement for oxidizing conditions to potentiate covalent adduct formation from the catechol suggests that this route of tamoxifen bioactivation may not be significant under normal circumstances in vivo, where reducing conditions prevail. Under conditions of oxidative stress, however, catechol formation may enable the subsequent generation of a reactive quinone, that could damage protein and/or DNA. Indeed Bolton and colleagues have recently shown that this species is cytotoxic and can form adducts with deoxyguanosine and deoxythymidine nucleosides (43). By contrast, the low level adduct formation observed with P450s 2B6 and 2C19 appears not to be dependent upon the concomitant production of ROS. Thus, adduct formation by these forms may be physiologically relevant. Though the low levels of binding seen here with these forms prevented an analysis of the targets of adduct formation, the fact that covalent binding was observed at subtherapeutic levels argues for the importance of this investigation.

Acknowledgment. Grateful thanks are extended to Drs. Judy Bolton, David Phillips, Martin Lennard, and Ian White for supplying metabolic standards used in this work, and to Professor K. Nishihara for the gift of the pGro7 vector. The authors sincerely appreciate helpful discussions with Drs. Ute Kent, Paul Hollenberg, Martin Lennard, David Phillips, and James DeVoss. Funding for this study was provided by the Kathleen Cuningham Foundation for Breast Cancer Research and the Australian Cancer Fund.

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TX0100439

Bioactivation of Tamoxifen by Recombinant Human Cytochrome P450

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