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Hexestrol (HES), a synthetic nonsteroidal estrogen, is carcinogenic in Syrian golden hamsters. ... double bond, are also carcinogenic in the kidney of...
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Chem. Res. Toxicol. 1998, 11, 412-419

Metabolic Activation and Formation of DNA Adducts of Hexestrol, a Synthetic Nonsteroidal Carcinogenic Estrogen Shyi-Tai Jan,† Prabu D. Devanesan,† Douglas E. Stack,† Ragulan Ramanathan,‡ Jaeman Byun,‡ Michael L. Gross,‡ Eleanor G. Rogan,† and Ercole L. Cavalieri*,† Eppley Institute for Research in Cancer and Allied Diseases, University of Nebraska Medical Center, 600 South 42nd Street, Omaha, Nebraska 68198-6805, and Department of Chemistry, Washington University, One Brookings Drive, St. Louis, Missouri 63130-4899 Received August 11, 1997

Hexestrol (HES), a synthetic nonsteroidal estrogen, is carcinogenic in Syrian golden hamsters. The major metabolite of HES is its catechol, 3′-OH-HES, which can be metabolically converted to the electrophilic catechol quinone, HES-3′,4′-Q, by peroxidases and cytochrome P450. Standard adducts were synthesized by reacting HES-3′,4′-Q with dG and dA to produce the adducts 3′-OH-HES-6′(R,β)-N7Gua and HES-3′,4′-Q-6′-N6dA, respectively. When HES-3′,4′-Q was reacted with calf thymus DNA, 3′-OH-HES-6′(R,β)-N7Gua was identified by HPLC and tandem mass spectrometry as the depurinating adduct, with minor amounts of stable adducts. 3′-OH-HES was bound to DNA after activation by horseradish peroxidase, lactoperoxidase, or rat liver microsomes. The depurinating adduct 3′-OH-HES-6′(R,β)-N7Gua was identified in these systems at levels of 65, 41, and 11 µmol/mol of DNA-P, respectively. Unidentified stable adducts were observed in much lower amounts and were quantified by the 32P-postlabeling method. Similarly to 3′-OH-HES, the catechol metabolites of the natural steroidal estrogens estrone (E1) and estradiol (E2), namely, 2-OHE1, 4-OHE1, 2-OHE2, and 4-OHE2, can be oxidized to their corresponding quinones by peroxidases and cytochrome P450. The quinones of the carcinogenic 4-OHE1 and 4-OHE2 have chemical and biochemical properties similar to those of HES-3′,4′-Q. The results suggest that formation of HES-3′,4′-Q may be a critical event in tumor initiation by HES and that HES is an excellent model compound to corroborate the hypothesis that estrogen-3,4-quinones are ultimate carcinogenic metabolites of the natural steroidal estrogens E1 and E2.

Introduction Comprehensive carcinogenicity studies on polycyclic aromatic hydrocarbons (PAH)1 (1, 2) indicate that depurinating PAH-DNA adducts, namely, the adducts lost from DNA by cleavage of the bond between deoxyribose and purine bases, generate the critical mutations found in the Harvey-ras oncogene of PAH-induced mouse skin papillomas (3). These mutations arise by misreplication of unrepaired apurinic sites produced by loss of depurinating adducts. This relationship led us to hypothesize that oxidation of 4-catechol estrogen metabolites of estrone (E1) and estradiol (E2), which are carcinogenic (4, 5), would result in 4-catechol estrogen quinones that also react with DNA to form depurinating adducts (6, 7). * To whom correspondence should be addressed. † University of Nebraska Medical Center. ‡ Washington University. 1 Abbreviations: CuOOH, cumene hydroperoxide; dA, deoxyadenosine; DES, diethylstilbestrol; dG, deoxyguanosine; DMF, dimethylformamide; ESI, electrospray ionization; E1, estrone; E2, estradiol; E13,4-Q, estrone-3,4-quinone; E2-3,4-Q, estradiol-3,4-quinone; FAB, fast atom bombardment; Gua, guanine; HES, hexestrol [3-(4′-dihydroxyphenyl)-4-(4"-hydroxyphenyl)hexane]; HES-3′,4′-Q, hexestrol-3′,4′quinone; HRP, horseradish peroxidase; IP, ionization potential(s); LP, lactoperoxidase; MS, mass spectrometry; 3′-OH-DES, 3′-hydroxydiethylstilbestrol; 2-OHE1, 2-hydroxyestrone; 4-OHE1, 4-hydroxyestrone; 2-OHE2, 2-hydroxyestradiol; 4-OHE2, 4-hydroxyestradiol; 3′-OH-HES, 3′-hydroxyhexestrol; PAH, polycyclic aromatic hydrocarbon(s); SCF, self-consistent field.

The nonsteroidal synthetic diethylstilbestrol (DES) and hexestrol, 3-(4′-dihydroxyphenyl)-4-(4"-hydroxyphenyl)hexane (HES), which is DES hydrogenated at the C-C double bond, are also carcinogenic in the kidney of Syrian golden hamsters (8, 9). The major metabolites of DES and HES are their catechols 3′-OH-DES and 3′-OH-HES (9-12). These catechols may be metabolically converted to catechol quinones. In this article, we report the reaction of HES-3′,4′quinone (HES-3′,4′-Q) with DNA to form depurinating adducts comparable to those observed with estrogen-3,4quinones (7). Furthermore, 3′-OH-HES was activated by horseradish peroxidase (HRP), lactoperoxidase (LP), or rat liver microsomal cytochrome P450 in the presence of DNA to form the same depurinating adducts. These depurinating adducts were identified and quantified by HPLC and tandem mass spectrometry (MS).

Experimental Procedures Caution. 3′-OH-HES and HES-3′,4′-Q may be carcinogenic and should be handled according to NIH guidelines (13). Chemicals. Deoxyadenosine (dA) and deoxyguanosine (dG) monohydrates were purchased from TCI (Portland, OR). HRP, LP, cumene hydroperoxide (CuOOH), and H2O2 were purchased from Sigma (St. Louis, MO). Other chemicals were purchased from Aldrich Chemical Co., WI). All chemicals were used without further purification.

S0893-228x(97)00141-0 CCC: $15.00 © 1998 American Chemical Society Published on Web 04/09/1998

Activation of Hexestrol To Form DNA Adducts Computational Methods. All calculations were carried out on a Silicon Graphics Iris Indigo 2 workstation using either SYBYL version 6.3 (Tripos Assoc., St. Louis, MO) or Spartan version 4.1 (Wavefunction, Inc., Irvine, CA) software. Rotational barriers were determined by using the grid search feature in SYBYL. Ionization potentials (IP) and atomic charges were calculated on Spartan by using the semiempirical Austin model 1 (AM1) calculation. Geometry optimizations were conducted by using the MMOK keyword for molecular mechanics correction to the guanine amide bond. In addition, the “converge” option and the “nopseudo” keyword were used to overcome poor selfconsistent field (SCF) convergence. HPLC. Preparative HPLC was conducted on a Waters (Milford, MA) 600E solvent delivery system equipped with a 484 tunable absorbance detector operating at 254 nm. Analytical HPLC was conducted on a Waters 600E solvent delivery system equipped with a 996 photodiode array detector interfaced to a NEC Powermate 486/33i computer. Analytical HPLC using electrochemical detection was conducted on a Scientific Systems (State College, PA) model 300 LC pump equipped with an ESA (Chelmsford, MA) Coulochem II electrochemical detector employing a porous carbon electrode set at 700 mV and monitored on a Hewlett-Packard (Kennett Square, PA) HP 3396A integrator. Under these conditions the limit of detection for catechol adducts is in the low-picomolar range. Analytical HPLC (reverse-phase) was conducted by using a YMC ODS-AQ 5-µm, 120-Å column (6.0 × 250 mm; YMC, Morris Plains, NJ). Preparative HPLC (reverse-phase) was conducted by using a YMC ODS-AQ 5-µm, 120-Å column (20 × 250 mm). NMR. 1H NMR spectra were recorded on samples in Me2SOd6 or acetone-d6 at room temperature on either a Varian XL 300-MHz or a Varian Unity 500-MHz (Palo Alto, CA) instrument, with chemical shifts reported relative to Me2SO-d6 at 2.49 ppm or to acetone-d6 at 2.04 ppm. Coupling constants (J) are given in hertz. Mass Spectrometry. Tandem mass spectra of fast atom bombardment (FAB)- and electrospray ionization (ESI)-produced ions and the mass spectra used for quantification were obtained with a VG ZAB-T four-sector mass spectrometer (Manchester, U.K.) of BEBE design (14). Exact mass measurements of FABproduced ions were obtained by peak matching with a Kratos MS-50 triple analyzer mass spectrometer (Manchester, U.K.) (15) at a mass resolving power of 10 000. Ions were desorbed from the probe tip by bombarding a mixture of sample and matrix with 6-7-keV argon atoms, and mass calibration was with a mixture of CsI and glycerol. (i) FAB Mass Spectrometry. Samples were dissolved in methanol, and a 1-µL aliquot was deposited on the probe tip in a matrix of 3-nitrobenzyl alcohol/glycerol (1:1). A primary beam of 22-keV cesium ions was used to desorb the analyte ions, which were accelerated to 8 kV. FAB tandem mass spectra were obtained by activating the precursor ion in a floated (4-kV) collision cell with helium at sufficient pressure to suppress the beam by 50%. The fragment ions were detected by using the final point detector. For ESI experiments, the accelerating potential was 4 kV, and the collision cell was floated at 2 kV. (ii) ESI Mass Spectrometry. The spray needle of the VG ESI source was maintained at 8000 V, the counter electrode (pepper pot) was at 5000 V, and the sampling cone, skimmer lens, skimmer, hexapole, and ring electrode were at 4177, 4125, 4119, 4117, and 4116 V, respectively. Nitrogen was used as both bath and nebulizer gas (at 80 °C) with a flow rate of approximately 300 and 12 L/h, respectively. A Harvard model 22 syringe pump (South Natick, MA) was used to infuse a solution of 50:49:1 methanol/water/acetic acid to the spray needle at a rate of 10 µL/min. To quantify the in vitro adducts, 5-µL aliquots of a 3′-OHHES-6′(R,β)-N7Gua reference sample with concentrations ranging from 25 fmol/µL to 5 pmol/µL were flow-injected via a sixport Rheodyne 7125 loop injector. Mass spectra of these standards and the unknowns were obtained by scanning over 60 Da at a rate of 35 s/decade (1-s delay between scans), and a

Chem. Res. Toxicol., Vol. 11, No. 5, 1998 413 calibration plot was made. For all experiments, 30 scans were signal-averaged, and an appropriate background subtract was done. A solution of a mixture of PEG 400 and 600 was used to calibrate the mass scale of the mass spectrometer. Synthesis of HES-3′,4′-Q and Deoxyribonucleoside Adducts. HES-3′,4′-Q. Ag2O (29 mg, 0.12 mmol) was added to a solution of 3′-OH-HES (12 mg, 0.04 mmol) (16) in acetone-d6 (1 mL) at room temperature. The reaction mixture was stirred at room temperature under nitrogen for 10 min and then filtered directly into a tube for NMR spectroscopy. The yield of this reaction was nearly quantitative: NMR (500 MHz, acetone-d6) 8.14 (s, 1H, Ar-OH), 7.17 (dd, J ) 10.5, 2.0 Hz, 1H, 6′-H), 6.98 (d, J ) 8.5 Hz, 2H, Ar-H), 6.70 (d, J ) 8.5 Hz, 2H, Ar-H), 6.28 (d, J ) 10.5 Hz, 1H, 5′-H), 6.17 (d, J ) 2.0 Hz, 1H, 2′-H), 2.47 (m, 2H, CH), 1.60 (m, 1H, CH2), 1.38 (m, 1H, CH2), 1.23 (m, 2H, CH2), 0.60 (dd, J ) 7.5, 7.5 Hz, 3H, CH3), 0.50 (dd, J ) 7.5, 7.5 Hz, 3H, CH3). Reaction of HES-3′,4′-Q with dG. The freshly prepared HES-3′,4′-Q (0.1 mmol) in 1 mL of acetone was filtered directly into a stirred solution of dG (285 mg, 1.0 mmol) in CH3CO2H/ H2O (4 mL, 1:1). After 6 h at room temperature, the solvent was removed under vacuum. The residue was dissolved in CH3OH/dimethylformamide (DMF) (1.5 mL, 1:1) and then isolated by preparative HPLC equipped with a UV detector at a flow rate of 6 mL/min. The column was eluted with CH3OH/ H2O (30:70) for 5 min, followed by a linear gradient (CV6) to 100% CH3OH in 70 min. 3′-OH-HES-6′(R,β)-N7Gua2 (retention time ) 48 min) was the only product isolated from this reaction in 80% yield: NMR (500 MHz, Me2SO-d6) 10.77 and 10.62 [bs, 1H, 1-NH(Gua), exchangeable with D2O], 9.14 (m, 3H, Ar-OH, exchangeable with D2O), 7.69 and 7.60 [s, 1H, 8-H(Gua)], 6.746.51 (m, 6H, Ar-H), 6.08 [bs, 2H, 2-NH2(Gua), exchangeable with D2O], 2.34 and 2.23 (m, 1H, CH), 2.05 and 1.96 (m, 1H, CH), 1.39 (m, 1H, CH2), 1.21 (m, 1H, CH2), 1.13 and 0.85 (m, 2H, CH2), 0.52 and 0.47 (dd, 3H, CH3), 0.31 (m, 3H, CH3 ); exact mass (M + H)+ calcd for C23H26O4N5 436.1985, obsd 436.1984. Reaction of HES-3′,4′-Q with dA. Freshly prepared HES3′,4′-Q (0.1 mmol) in 1 mL of acetone was filtered directly into a stirred solution of dA (269 mg, 1.0 mmol) in CH3CO2H/H2O (4 mL, 1:1). After 6 h at room temperature, the solvents were removed under vacuum. The residue was dissolved in CH3OH/ DMF (1.5 mL, 1:1) and then isolated by two consecutive preparative HPLC runs to afford HES-3′,4′-Q-6′-N6dA with a yield of 31%. The first HPLC run was conducted as described for the separation of the dG adduct. The N6dA adduct coeluted with an impurity at a retention time of 55 min. The second HPLC run was conducted by using an isocratic condition of CH3CN/H2O (40:60) at a flow rate of 6 mL/min, and the retention time of the adduct was 28 min: NMR (500 MHz, Me2SO-d6) 9.17 and 8.63 [s and s, 2H, 2-H and 8-H(dA)], 8.31 (s, 1H, Ar-OH, exchangeable with D2O), 7.04 (m, 3H, Ar-H), 6.83 (s, 1H, ArH), 6.74 (d, J ) 8.5 Hz, 2H, Ar-H), 6.59 [bs, 1H, 6-NH(dA), exchangeable with D2O], 6.51 [dd, J ) 6.0, 6.0 Hz, 1H, 1′-H(dA)], 5.39 [bs, 1H, 3′-OH(dA), exchangeable with D2O], 5.00 [bs, 1H, 5′-OH(dA), exchangeable with D2O], 4.44 [m, 1H, 3′-H(dA)], 3.90 [m, 1H, 4′-H(dA)], 3.61 [m, 1H, 5′-H(dA)], 3.56 [m, 1H, 5′-H(dA)], 2.86 [m, 1H, 2′-H(dA)], 2.73 [m, 1H, 2′-H(dA)], 2.42 (m, 2H, CH), 1.33-1.23 (m, 4H, CH2), 0.43 (m, 6H, CH3); exact mass (M + H)+ calcd for C28H32O6N5 534.2353, obsd 534.2352. Covalent Binding of Quinones to DNA. HES-3′,4′-Q (1 mg/50 µL of Me2SO) was mixed with 6 mM calf thymus DNA in 0.067 M sodium-potassium phosphate (pH 7.0) and incubated for 2 h at 37 °C. The solution was extracted twice with CHCl3, DNA was precipitated with two volumes of ethanol, and the supernatant was used for analysis of depurinating adducts. The DNA was redissolved in 0.015 M NaCl-0.0015 M sodium citrate, the concentration was determined by its absorbance at 2 The terminology (R,β) is used to indicate that the adduct has two rotational conformers that can be seen by NMR but are not separable. The R and β isomers have the purine base below or above the plane of HES, respectively.

414 Chem. Res. Toxicol., Vol. 11, No. 5, 1998 260 nm, and the DNA was used for analysis of stable adducts by the 32P-postlabeling method (17). Covalent Binding of 3′-OH-HES to DNA. 3′-OH-HES was bound to DNA in reactions catalyzed by HRP or LP in the presence of H2O2 or with phenobarbital-induced rat liver microsomal cytochrome P450 in the presence of CuOOH. In the 15-mL peroxidase-catalyzed reactions, mixtures containing 3 mM calf thymus DNA in 0.067 M sodium-potassium phosphate (pH 7.0), 3′-OH-HES (1 mg/50 µL of Me2SO), 0.5 mM H2O2, and 100 µg/mL HRP (31 units, type VI) or 100 µg/mL LP (9 units) were incubated for 2 h at 37 °C. H2O2 was added at 30-min intervals. For the microsome-catalyzed reaction, 15-mL mixtures containing 3 mM calf thymus DNA in 150 mM Tris-HCl, pH 7.5, 150 mM KCl, 5 mM MgCl2, 3′-OH-HES (1 mg/50 µL of Me2SO), 15 mg of microsomal protein, 0.6 mM NADPH or 1 mM CuOOH were incubated for 2 h at 37 °C. A 1-mL aliquot was used for analysis of stable DNA adducts after purification of the DNA by extraction with phenol and CHCl3 and precipitation with ethanol, as previously reported (17). DNA was precipitated from the rest of the incubation mixture with two volumes of ethanol, and the supernatant was used for analysis of depurinating adducts. Control reactions for each type of enzyme-catalyzed DNAbinding reaction were carried out under identical conditions with either no enzyme or no cofactor (NADPH, H2O2, or CuOOH). Identification of Depurinating Adducts. The supernatant from each in vitro reaction, containing depurinating adducts and metabolites, was dried under vacuum. The solid material was dissolved in CH3OH/DMF (1.5 mL, 1:1) and filtered to remove undissolved material prior to initial analysis by preparative HPLC. A sample of the standard, 3′-OH-HES6′(R,β)-N7Gua, was first injected into the preparative HPLC to determine its exact retention time (ca. 48 min) by UV detection. The column was initially eluted with CH3OH/H2O (30:70) for 5 min, followed by a linear gradient to 100% CH3OH in 70 min at a flow rate of 6 mL/min. The biological samples were then injected under the same conditions, and the fraction that had the retention time of the standard was collected and dried under vacuum for the second HPLC analysis, which was conducted by using an analytical HPLC equipped with an electrochemical detector. The column was eluted with CH3CN/0.03 M ammonium phosphate, pH 3.5 (1.2:3), under isocratic conditions at a flow rate of 0.9 mL/min. A known amount of the standard adduct, 3′-OH-HES-6′(R,β)-N7Gua, was first injected to obtain its retention time, typically 15 min, as well as its peak intensity. The biological sample collected in the first HPLC was dissolved in 1 mL of CH3OH, and 20 µL of this solution was injected into the analytical HPLC. The peak eluting at the retention time of the standard adduct was assigned as 3′-OH-HES-6′(R,β)N7Gua. Assignment of the 3′-OH-HES-6′(R,β)-N7Gua peak was further confirmed by coinjection of the standard adduct and biological sample and by mass spectrometry. The amount of biologically formed depurinating adduct eluted from the analytical HPLC was determined by the area of the electrochemical signal in relation to the signal generated by a known amount of standard adduct. Analysis of Stable Adducts by 32P-Postlabeling. 32PPostlabeling analysis for the determination of stable DNA adducts was carried out with 8 µg of DNA on 10- × 13-cm PEIcellulose TLC plates by using the chromatographic conditions and solvents published previously (17). The adduct spots were detected by autoradiography, excised, and counted for radioactivity with a liquid scintillation counter. No adduct spots were observed for samples from the control reactions.

Results and Discussion Synthesis and Structure Elucidation of HES3′,4′-Q Adducts Formed with dG and dA. Reaction of HES-3′,4′-Q with dG and dA, followed by structure determination of the resulting adducts, revealed the electrophilic properties of HES-3′,4′-Q, the site of covalent

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binding with dG and dA, and their chromatographic and mass spectral properties. These synthesized adducts serve as markers for analyzing biologically formed DNA adducts. 3′-OH-HES was quantitatively converted to HES3′,4′-Q by oxidation with Ag2O. After 6 h at room temperature, reaction of HES-3′,4′-Q with dG yielded 3′OH-HES-6′(R,β)-N7Gua (80% yield), which eluted as a single peak under different HPLC conditions (Scheme 1). In contrast to the dG adduct, the dA adduct was isolated as a quinone derivative. Under the same conditions used for dG, reaction of HES-3′,4′-Q with dA yielded HES-3′,4′Q-6′-N6dA as the only adduct, with a yield of 31% (Scheme 1). HES-3′,4′-Q did not react with deoxycytidine or thymidine. 3′-OH-HES-6′(r,β)-N7Gua. The NMR spectrum for the 3′-OH-HES-6′(R,β)-N7Gua adduct shows two sets of resonance signals (Figure 1) that were initially attributed to the formation of two regioisomers via attack of dG at the 5′- and 6′-positions. A better explanation, as discussed below, is the existence of two atropisomers (rotational isomers) that result from hindered rotation about the C6′-N7Gua bond. The NMR chemical shifts and integration of all of the aliphatic protons in the adduct, namely, the two ethyl groups and two benzylic protons, indicate that all of the sp3 protons are unaffected by adduction (Figure 1). A two-dimensional chemical shift correlation spectroscopy spectrum (not shown) was taken to confirm these aliphatic protons. The loss of the deoxyribose moiety from the adduct and the presence of the chemical shifts assigned to 1-NH, 8-H, and 2-NH2 suggest that the 3′OH-HES moiety was bound at the N7 of Gua. The NMR spectrum also indicated that 1-NH(Gua) and 8-H(Gua) had two distinct sets of chemical shifts at 10.77/10.62 and 7.69/7.60 ppm, respectively. Two sets of the benzylic proton resonances also showed typical chemical shifts of these benzylic protons at 2.34, 2.23, 2.05, and 1.96 ppm. The three phenolic protons of each adduct gave a multiplet at 9.14 ppm, which is similar to the chemical shift of the phenolic protons of 3′-OH-HES, indicating that the quinone has reverted to catechol in the formation of the adduct. This is corroborated by the mass spectrum of this adduct (see below). The chemical shifts of all aromatic protons appear in the narrow region between 6.74 and 6.51 ppm. The rationale that the two isomers shown by NMR are rotational isomers at C6′, and not positional isomers, is based on the similar chemistry of the estrogen-3,4quinones (6). In fact, reaction with estrogen-3,4-quinones has demonstrated that nitrogen nucleophiles, under both neutral and acidic conditions, undergo Michael addition to the unsubstituted carbon, C1, which bears the greatest positive charge. “Hard” nucleophiles, such as nitrogen and oxygen, attack estrogen quinones exclusively at the 1-position (6, 18) (analogous to the 6′-position of 3′-OHHES), whereas “soft” nucleophiles, such as sulfur, attack the 2-position (19) [analogous to the 5′-position of 3′-OHHES (9)]. In fact, Mulliken charge calculations, derived from semiempirical quantum mechanical calculations using the AM1 Hamiltonian, show the 6′-position to have the highest positive charge character of all unsubstituted carbons (Scheme 1). Thus, the presence of two sets of NMR signals in the 3′-OH-HES-6′(R,β)-N7Gua spectrum is attributed to formation of atropisomers, which exist owing to restricted rotation about the N7Gua-C6′ bond.

Activation of Hexestrol To Form DNA Adducts

Chem. Res. Toxicol., Vol. 11, No. 5, 1998 415

Scheme 1. Oxidation of 3′-OH-HES to HES-3′,4′-Q and Reaction of the Quinone with dG and dAa

a

Charge distribution in HES-3′,4′-Q, ionization potentials, and rotational energy are shown.

Figure 1. NMR spectrum of 3′-OH-HES-6′(R,β)-N7Gua.

This same restriction of rotation was observed in estrogen quinone adducts of guanine in which its N7 is bonded to the C1 of the estrogen (6). The more sterically rigid estrogen ring system produces a very high rotational barrier of ca. 600 kcal/mol (6). The 3′-OH-HES-6′(R,β)-N7Gua adduct (Scheme 1) has a 28.4 kcal/mol rotational barrier (see below). This permits the two sets of NMR signals observed at room

temperature (Figure 1). The rotational barrier results because the C8 proton of guanine approaches the van der Waals radius of the C2 methylene protons of the HES moiety as the guanine ring rotates about the N7-C6′ bond. The barrier was calculated by first rotating all bonds affecting the proximity of the C8(Gua) and C2(HES) protons (molecular mechanics, Tripos force field). The rotation of the C1-C2, C2-C3, C3-C4, and C3-C1′

416 Chem. Res. Toxicol., Vol. 11, No. 5, 1998

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Figure 2. Product-ion spectrum from collisionally activated decomposition of 3′-OH-HES-6′(R,β)-N7Gua introduced as [M + H]+ by ESI: (A) spectrum of synthetic reference; (B) spectrum of adduct isolated from in vitro study.

bonds generated more than 600 conformers. Two criteria were used to narrow the number of conformers in the second step of the rotational barrier search, namely, rotation about the N7-C6′ bond. Structures were selected that had a minimum C8(Gua) proton-to-C2(HES) proton distance of 5.2 Å (allowing for a maximum probability of rotation) and a molecular mechanics energy of no more than 11.0 kcal/mol above the global minimum. A total of 17 low-energy conformers were produced. They were subjected to a torsional driver, rotating the N7C6′ bond. The lowest energy barrier for any of the conformers was found to be 28.4 kcal/mol. This barrier is sufficient to produce two sets of resonance signals on the NMR time scale. To isolate two species in equilibrium (20), an energy barrier of ca. 23 kcal/mol is necessary. Rotational isomers can be observed by NMR spectroscopy, and the free energy of rotation (∆Grot) can be calculated if the rotational barrier occurs within the NMR time scale at temperatures conducive to NMR instrumentation (-100 to 150 °C). The relationship between the coalescence temperature, ∆Grot, and the difference in resonance signals of two protons made unequivalent due to restricted rotation is given by the Eyring equation.3 Using this equation and the difference in chemical shifts for the C8 protons (∆ppm ) 0.09, kr ) 45 Hz at 500 MHz), the ∆Grot that affords coalescence at 298 K is 14.6 kcal/mol. Thus, two sets of NMR resonance signals from rotational isomers would be observed only if ∆Grot is greater than ca. 15 kcal/mol. The rotational barrier for 3′-OH-HES-6′(R,β)-N7Gua is ca. 28 kcal/mol. At this ∆Grot, a coalescence temperature needed to merge the two C8 signals would be 555 K (282 °C), well above the limit for NMR investigation. Mass spectrometry provided additional structural information. Exact mass measurement of the FABproduced [M + H]+ was within 0.0001 u of the calculated value. Structural information was obtained by tandem 3 Eyring equation: ∆G rot (J/mol) ) 19.1Tc[10.32 + log(Tc/kr)], where Tc is the temperature of coalescence (K) and kr is the exchange frequency (Hz) (21).

mass spectrometry of [M + H]+ ions that were produced by both FAB and ESI (Figure 2A). The principal product ions formed upon high-energy collisional activation are of m/z 300 and 286. The former ion results from a chargeremote cleavage of the C3-C4 bond of the HES with elimination of H2 and β-methyl-p-hydroxystyrene (C9H10O). A related reaction is a simple cleavage of the C3-C4 bond to produce presumably a distonic radical cation of m/z 301. The m/z 286 ion is formed by an analogous chargeremote process involving loss of methane and the same C9H10O. These three processes offer additional evidence that the base is substituted on the catechol ring. There are a few low-abundance product ions that deserve discussion. The ions of m/z 406 and 378 arise by charge-remote losses of the elements C2H6 followed by loss of C2H4, consistent with the two ethyl groups. The relative abundances of these ions are more pronounced (by a factor of 2) when the precursor ion is produced by FAB. FAB-produced ions also yielded low-abundance (99%) by E1-3,4-Q or E2-3,4-Q is the formation of depurinating adducts (7). The stable adducts are formed in much lower amounts, less than 1% of the total.

Conclusions The catechol of HES, which is its major metabolite (9, 12), can be further oxidized to HES-3′,4′-Q by peroxidases and cytochrome P450. The quinone reacts with DNA to form predominantly the depurinating adduct 3′-OH-HES6′(R,β)-N7Gua and much lower levels of unidentified stable adducts. Comparison of these results with those obtained in a study of the 4-catechol estrogens of E1 and E2, shows that high levels of comparable N7Gua adducts are formed in that system (6, 7). In contrast, the 2-catechol estrogens of E1 and E2 form only stable adducts (6, 17). E1, E2 and their 4-catechol estrogens are carcinogenic in the kidney of Syrian golden hamsters, whereas the 2-catechol estrogens are not (4, 5). Because HES is carcinogenic in the hamster kidney (8, 9) and oxidation of its catechol in the presence of DNA leads to formation of N7Gua depurinating adducts, we suggest that formation of DNA adducts by HES-3′,4′-Q represents a critical event in tumor initiation by HES. Furthermore, these results are consistent with the hypothesis that estrogen-3,4-quinones are endogenous tumor initiators.

Acknowledgment. Dr. Jan was supported by a postdoctoral fellowship from the Eppley Institute Cancer Research Training Program Grant T32 CA09476. This research was supported by a U.S. PHS grant from the National Cancer Institute (P01 CA49210) and a grant from the Nebraska Department of Health. Core support at the Eppley Institute was funded by NCI Laboratory Cancer Research Center Support (Core) Grant CA36727. The mass spectrometry research resource at Washington University is supported by the NIH (2P41RR00954).

References (1) Cavalieri, E. L., and Rogan, E. G. (1992) The approach to understanding aromatic hydrocarbon carcinogenesis. The central role of radical cations in metabolic activation. Pharmacol. Ther. 55, 183-199. (2) Cavalieri, E., and Rogan, E. (1998) Mechanisms of tumor initiation by polycyclic aromatic hydrocarbons in mammals. In The Handbook of Environmental Chemistry, Vol. 3J: PAHs and Related Compounds (Neilson, A. H., Ed.) pp 81-117, SpringerVerlag, Heidelberg. (3) Chakravarti, D., Pelling, J. C., Cavalieri, E. L., and Rogan, E. G. (1995) Relating aromatic hydrocarbon-induced DNA adducts and c-Harvey-ras mutations in mouse skin papillomas: The role of apurinic sites. Proc. Natl. Acad. Sci. U.S.A. 92, 10422-10426.

Chem. Res. Toxicol., Vol. 11, No. 5, 1998 419 (4) Liehr, J. G., Fang, W. F., Sirbasku, D. A., and Ari-Ulubelen, A. (1986) Carcinogenicity of catechol estrogens in Syrian hamsters. J. Steroid Biochem. 24, 353-356. (5) Li, J. J., and Li, S. A. (1987) Estrogen carcinogenesis in Syrian hamster tissues: Role of metabolism. Fed. Proc. 46, 1858-1863. (6) Stack, D. E., Byun, J., Gross, M. L., Rogan, E. G., and Cavalieri, E. (1996) Molecular characteristics of catechol estrogen quinones in reactions with deoxyribonucleosides. Chem. Res. Toxicol. 9, 851-859. (7) Cavalieri, E. L., Stack, D. E., Devanesan, P. D., Todorovic, R., Dwivedy, I., Higginbotham, S., Johansson, S. L., Patil, K. D., Gross, M. L., Gooden, J. K., Ramanathan, R., Cerny, R. L., and Rogan, E. G. (1997) Molecular origin of cancer: Catechol estrogen3,4-quinones as endogenous tumor initiators. Proc. Natl. Acad. Sci. U.S.A. 94, 10937-10942. (8) Li, J. J., Li, S. A., Klicka, J. K., Parsons, J. A., and Lam, L. K. T. (1983) Relative carcinogenic activity of various synthetic and natural estrogens in the Syrian hamster kidney. Cancer Res. 43, 5200-5204. (9) Liehr, J. G., Ballatore, A. M., Dague, B. B., and Ulubelen, A. A. (1985) Carcinogenicity and metabolic activation of hexestrol. Chem.-Biol. Interact. 55, 157-176. (10) Haaf, H., and Metzler, M. (1985) In vitro metabolism of diethylstilbestrol by hepatic, renal and uterine microsomes of rats and hamsters. Biochem. Pharmacol. 34, 3107-3115. (11) Blaich, G., Gåttlicher, M., Cikryt, P., and Metzler, M. (1990) Effects of various inducers on diethylstilbestrol metabolism, drugmetabolizing enzyme activities and the aromatic hydrocarbon (Ah) receptor in male Syrian golden hamster liver. J. Steroid Biochem. 35, 201-204. (12) Metzler, M., and McLachlan, J. A. (1981) Oxidative metabolism of the synthetic estrogens hexestrol and dienestrol indicates reactive intermediates. Adv. Exp. Med. Biol. 136A, 829-837. (13) NIH Guidelines for the Laboratory Use of Chemical Carcinogens (1981), NIH Publications No. 81-2385, U.S. Government Printing Office, Washington, D.C. (14) Gross, M. L. (1990) Tandem mass spectrometry: Multisector magnetic instruments. In Methods in Enzymology, Vol. 193: Mass Spectrometry (McCloskey, J. A., Ed.) pp 131-153, Academic Press, San Diego, CA. (15) Gross, M. L., Chess, E. K., Lyon, P. A., Crow, F. W., Evans, S., and Tudge, H. (1982) Triple analyzer mass spectrometry for highresolution MS/MS studies. Int. J. Mass Spectrom. Ion Phys. 42, 243-254. (16) Jan, S.-T., Rogan, E. G., and Cavalieri, E. L. (1998) Large-scale synthesis of the catechol metabolites of diethylstilbestrol and hexestrol. Chem. Res. Toxicol. 11, 408-411. (17) Dwivedy, I., Devanesan, P. D., Cremonesi, P., Rogan, E. G., and Cavalieri, E. L. (1992) Synthesis and characterization of estrogen 2,3- and 3,4-quinones. Comparison of the DNA adducts formed by the quinones versus horseradish peroxidase-activated catechol estrogens. Chem. Res. Toxicol. 5, 828-833. (18) Abul-Hajj, Y. J., Tabakovic, K., Gleason, W. B., and Ojala, W. H. (1996) Reactions of 3,4-estrone quinone with mimics of amino acid side chains. Chem. Res. Toxicol. 9, 434-438. (19) Abul-Hajj, Y. J., and Cisek, P. L. (1988) Catechol estrogen adducts. J. Steroid Biochem. 31, 107-110. (20) Kalinowski, H. O., and Kessler, H. (1973) Fast isomerizations about double bonds. Top. Stereochem. 7, 295-383. (21) Breitmaier, E. (1993) In Structure Elucidation by NMR in Organic Chemistry (Breitmaier, E., Ed.) pp 61-63, Wiley, West Sussex, U.K. (22) Byun, J., Gooden, J., Ramanathan, R., Li, K.-M., Cavalieri, E., and Gross, M. L. (1997) Determination of isomeric dibenzo[a,l]pyrene-adenine adducts by six different tandem mass spectrometric experiments. J. Am. Soc. Mass Spectrom. 9, 977-986. (23) Kim, H. S., Yu, M., Jiang, Q., and LeBreton, P. (1993) UV photoelectron and ab initio quantum mechanical characterization of 2′-deoxyguanosine 5′-phosphate: Electronic influences on DNA alkylation patterns. J. Am. Chem. Soc. 115, 6169-6183. (24) Klopman, G. (1974) In Chemical Reactivity and Reaction Paths (Klopman, G., Ed.) pp 55-165, Wiley-Interscience, New York.

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