Synthesis and Structure Determination of the Adducts Formed by

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Articles Synthesis and Structure Determination of the Adducts Formed by Electrochemical Oxidation of Dibenzo[a,l]pyrene in the Presence of Adenine Kai-Ming Li,† Jaeman Byun,‡ Michael L. Gross,‡ Dan Zamzow,§ Ryszard Jankowiak,§ Eleanor G. Rogan,† and Ercole L. Cavalieri*,† Eppley Institute for Research in Cancer and Allied Diseases, University of Nebraska Medical Center, 986805 Nebraska Medical Center, Omaha, Nebraska 68198-6805, Department of Chemistry and Ames Laboratory-U.S. Department of Energy, Iowa State University, Ames, Iowa 50011, and Department of Chemistry, Washington University, One Brookings Drive, St. Louis, Missouri 63130 Received August 18, 1998

Because the radical cations of polycyclic aromatic hydrocarbons (PAH) are involved in tumor initiation, determination of the structures of biologically formed PAH-DNA adducts is important and relies on comparison of their properties with those of synthesized adducts. One of the possible sites of adduct formation is the N-3 position of Ade, but this depurinating adduct is not obtained by one-electron oxidation of dibenzo[a,l]pyrene (DB[a,l]P) in the presence of deoxyadenosine. Therefore, we turned to electrochemical oxidation of DB[a,l]P in the presence of Ade in dimethylformamide and produced the following adducts: DB[a,l]P-10-N1Ade (47%), DB[a,l]P-10-N3Ade (5%), DB[a,l]P-10-N7Ade (2%), and DB[a,l]P-10-N6Ade (6%). In Me2SO, this reaction afforded the same four adducts, but in slightly different yields: DB[a,l]P-10N1Ade (44%), DB[a,l]P-10-N3Ade (9%), DB[a,l]P-10-N7Ade (1%), and DB[a,l]P-10-N6Ade (3%). These adducts were purified by reverse-phase HPLC, and the subtle differences between the isomers were revealed by NMR, tandem mass spectrometry, and fluorescence line-narrowing spectroscopy. The relative yields of the N1Ade, N3Ade, and N7Ade adducts reflect the nucleophilicity and steric accessibility of these three nitrogen atoms in Ade.

Introduction Dibenzo[a,l]pyrene (DB[a,l]P)1 is the most potent carcinogenic polycyclic aromatic hydrocarbon (PAH) (1-5). Metabolic activation of DB[a,l]P leading to tumor initiation occurs via two main pathways: one-electron oxidation to yield the DB[a,l]P radical cation and monooxygenation to produce fjord-region diol epoxides (DB[a,l]PDE) (6, 7).2 Identification and quantification of DB[a,l]P-DNA adducts in biological systems not only can demonstrate that DB[a,l]P is activated by these two pathways but also can reveal the relative amounts of adducts formed in the different systems. * To whom correspondence should be addressed. † University of Nebraska Medical Center. ‡ Washington University. § Iowa State University. 1 Abbreviations: BP, benzo[a]pyrene; 1-CH Ade, 1-methyladenine; 3 COSY, homonuclear two-dimensional chemical shift correlation spectroscopy; dA, deoxyadenosine; DB[a,l]P, dibenzo[a,l]pyrene; DB[a,l]PDE, dibenzo[a,l]pyrene diol epoxide(s); dG, deoxyguanosine; DMF, dimethylformamide; ESI, electrospray ionization; FLNS, fluorescence line-narrowing spectroscopy; MS/MS, tandem mass spectrometry; NOE, nuclear Overhauser effect; PAH, polycyclic aromatic hydrocarbon(s). 2 K.-M. Li et al., Identification and quantitation of depurinating DNA adducts of dibenzo[a,l]pyrene formed in vitro and in mouse skin and rat mammary gland, to be submitted for publication.

To identify the biologically formed DB[a,l]P-DNA adducts, it is necessary to obtain prospective model adducts that will serve as reference standards. We synthesized standard adducts from DB[a,l]PDE with deoxyadenosine (dA) and deoxyguanosine (dG) (8, 9). One-electron oxidation model adducts were previously prepared by electrochemical oxidation of DB[a,l]P in the presence of dA and dG (10). We found, however, that by electrochemical oxidation of PAH in the presence of dA, the depurinating adducts with the PAH bonded to the N-3 of Ade are not obtained, owing to interference by the deoxyribose linked at the adjacent N-9 position of Ade (10-15). The N3Ade adducts of several PAHs are formed biologically, including those of benzo[a]pyrene (BP) (16), 7H-dibenzo[c,g]carbazole (17), and DB[a,l]P (6).2 Because it is generally possible to synthesize N3Ade adducts by one-electron oxidation of PAH in the presence of Ade (1417), we embarked on the synthesis of various DB[a,l]PAde adducts. In this article, we report the synthesis of DB[a,l]P-10N1Ade, DB[a,l]P-10-N3Ade, DB[a,l]P-10-N7Ade, and DB[a,l]P-10-N6Ade by electrochemical oxidation of DB[a,l]P in the presence of Ade. The peculiar properties of the N1Ade adducts of DB[a,l]P and other PAH in reversephase HPLC with CH3OH/H2O and CH3CN/H2O gradi-

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ents, which impeded detection of these adducts, were circumvented by using C2H5OH/H2O gradients. Because the DB[a,l]P-Ade isomers obtained by electrochemical oxidation are very similar, the combination of NMR, tandem mass spectrometry (MS/MS), and fluoresence line-narrowing spectroscopy (FLNS) reveals that subtle differences between the isomers can be identified, allowing ultimately their characterization in biological systems.

Experimental Methods Caution: DB[a,l]P is a hazardous chemical and was handled according to NIH guidelines (18). General Procedures. (1) HPLC. HPLC was conducted on a Waters 600E solvent delivery system equipped with a Waters 700 WISP autoinjector. Eluents were monitored for UV absorbance (254 nm) with a Waters 996 photodiode array detector, and data were collected on an APC-IV Powermate computer. Analytical runs were conducted by using a YMC ODS-AQ 5 µm, 120 Å column (6.0 mm × 250 mm) (YMC, Wilmington, NC). Two different HPLC conditions were used for adduct purification. One was a CH3CN/H2O gradient consisting of elution for 5 min with 20% CH3CN in H2O, followed by an 80 min linear gradient (CV6) to 100% CH3CN, carried out at a rate of 1.0 mL/min; the second condition consisted of elution for 5 min with 60% CH3CN/C2H5OH (3:1, v/v) in H2O, followed by a 40 min linear gradient to 100% CH3CN/C2H5OH (3:1, v/v). Preparative HPLC was conducted by using a YMC ODS-AQ 5 µm, 120 Å column (20 mm × 250 mm) at a flow rate of 6.0 mL/min. (2) UV. UV absorbance spectra were recorded by using a Waters 996 photodiode array detector during elution from HPLC with the CH3CN/H2O gradient described above, and elution was monitored at 308 nm. Yields of the adducts were determined by integration of the peak areas of the adducts at 308 nm, since the molar extinction coefficients of these adducts are very similar. (3) NMR. Proton and homonuclear two-dimensional chemical shift correlation spectroscopy (COSY) NMR spectra were recorded in Me2SO-d6 at 25 °C on a Varian Unity 500 NMR spectrometer (499.843 MHz). Chemical shifts were determined relative to the signal of Me2SO at 2.50 ppm. Nuclear Overhauser effect (NOE) difference spectra were recorded by applying a presaturation pulse with a decoupler on resonance and then subtracting the NMR trace from the corresponding reference spectrum recorded under identical conditions, but with the decoupler off-resonance. Typical spectra were acquired from at least 3000 transients. (4) Electrospray and Tandem Mass Spectrometry. Exactmass measurements were taken for [M + H]+ ions that were desorbed by fast atom bombardment (6-7 keV neon atoms from an Ion Tech FAB gun) into a Kratos MS-50 triple analyzer mass spectrometer (19). The resolving power of the first stage of the instrument was set at 10 000 (10% peak maximum), and the acceleration potential was 8000 V. The sample (approximately 10 nmol) was dissolved in 10 µL of CH3OH or Me2SO, and a 1 µL aliquot was loaded on the probe along with 1 µL of 3-nitrobenzyl alcohol and glycerol (1:1), which contained a saturated amount of CsI. Exact masses were determined by peak matching, using a nearby glycerol cluster or CsI ions as the mass standards. The measurements were within 1 ppm of the theoretical value. Low-energy collisionally activated decomposition spectra were acquired with Finnigan TSQ 7000 (San Jose, CA) and a Micromass Quattro II (Manchester, U.K.) mass spectrometers in the application/demonstration laboratories of the two manufacturers. All samples were introduced as a solution with a concentration of approximately 1 pmol/µL via an electrospray ion source at flow rates of 5 µL/min (TSQ 7000) and 2 µL/min (Quattro II). The electrospray needle was held at 4000 V and the counter electrode at ground potential. The solvent system,

Li et al. CH3OH/H2O/CH3CN/CH3COOH (1:1:1:0.03), was used to dissolve the samples and to serve as the carrier solvent for the electrospray source. To obtain product ion spectra in the MS/ MS mode, argon (TSQ 7000) and nitrogen (Quattro II) were used as collision gases. Product ion spectra were obtained at a collision energy of 55 eV (offset of Q2 was 55 V with respect to the voltage of Q1) by signal averaging 10-20 0.8 s scans. (5) FLNS. The instrumentation used for low-temperature, laser-excited fluorescence spectroscopy is described elsewhere (20); therefore, only the most important aspects are summarized here. The excitation source was a Lambda Physik (Acton, MA) Lextra XeCl excimer laser-FL-2002 dye laser system. FLN spectra were acquired with the excimer-pumped dye laser under FLN conditions (4.2 K, S1 r S0 excitation). Selective excitation at a variety of wavelengths provided unique FLN spectra for PAH adducts (20). Fluorescence was collected at a right angle to the laser excitation beam and dispersed by a McPherson (Acton, MA) model 2061 1 m monochromator. All fluorescence spectra were obtained by using a Princeton Instruments (Trenton, NJ) IRY-1024/GRB intensified diode array detector in gated detection mode, using the output of a reference photodiode to trigger a Princeton Instruments FG-100 high-voltage pulse generator. The FLN spectra were acquired using a 40 ns delay and a 200 ns gate width. Samples were dissolved in C2H5OH prior to analysis. Materials. DB[a,l]P was obtained from the National Cancer Institute Chemical Carcinogen Repository (Bethesda, MD). It was more than 99% pure as determined by HPLC (mp 161162 °C) and was used as received. Ade was purchased from Aldrich (Milwaukee, WI) and dA from TCI America (Portland, OR), and these compounds were desiccated over P2O5 under vacuum at 110 °C for 48 h prior to being used. Commercially available dimethylformamide (DMF) (Aldrich) was purified by heating to reflux over CaH2, followed by vacuum distillation just prior to use, and was stored over 4 Å molecular sieves under argon. KClO4 and 1-methyladenine (1-CH3Ade) were purchased from Aldrich and used as received. Electrochemical Synthesis of Adducts. Electrochemical synthesis was conducted as previously described (11). The oxidation potential used for the synthesis of DB[a,l]P adducts was 1.10 V, slightly less than its anodic peak potential of 1.14 V, measured by cyclic voltammetry (model CV27, Bioanalytical Systems, Lafayette, IN) in DMF. The oxidation potential of Ade was not determined in DMF because this solvent began to be oxidized at 1.45 V, indicating that the anodic potential of Ade is above this value. Thus, during adduct synthesis at 1.10 V, Ade was not oxidized. The electrochemical cell, electrodes, glassware, needles, and syringes were dried at 150 °C prior to being used. The electrochemical cell and the electrodes were assembled while hot and allowed to cool under argon. Coupling between DB[a,l]P and the nucleophilic groups of Ade or dA was accomplished by selective anodic oxidation of DB[a,l]P in the presence of Ade (Scheme 1) or dA (Scheme 2). To a solution of KClO4 (0.5 M) in DMF or Me2SO was applied an oxidation potential of 1.40 V for 30 min under argon. To the pre-electrolyzed solution were added DB[a,l]P (10 mg, 0.033 mmol) and Ade (44.7 mg, 0.33 mmol) or dA (83 mg, 0.33 mmol) in the electrochemical cell while it was turned off. After DB[a,l]P and Ade or dA were dissolved, the cell potential was again applied; the electrode potential was gradually raised from 0 to 1.10 V and kept constant at this value during the entire electrolysis. Both the output current and the total charge were monitored throughout the experiment. The reaction was stopped when the current had decreased to 1/20 of the initial value and a charge of 6.4 C (for a two-electron transfer) had accumulated. These two conditions were achieved in approximately 2 h. After the reaction was complete, DMF was removed under vacuum, the adducts were extracted four times from the residue by using a solvent mixture of C2H5OH/CHCl3/(CH3)2CO (2:1:1, v/v), and the combined extract was filtered through Whatman fluted filter paper. The filtrate was evaporated under vacuum,

Reaction of DB[a,l]P Radical Cation with Ade

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Scheme 1. Electrochemical Oxidation of DB[a,l]P in the Presence of Ade in DMF

Scheme 2. Electrochemical Oxidation of DB[a,l]P in the Presence of dA in DMF

and the residue was dissolved in 3 mL of Me2SO/CH3OH (1:1, v/v), filtered through a 0.45 mm filter, and analyzed by HPLC with the CH3CN/H2O and CH3CN/C2H5OH/H2O gradients. When the electrochemical oxidation was conducted in Me2SO, H2O was added at the end of the reaction and the adducts were extracted into CHCl3. Evaporation of the organic extract afforded a mixture of adducts. Purification of the adducts was conducted by using preparative HPLC with the CH3CN/H2O gradient, followed by the CH3CN/C2H5OH/H2O gradient.

The adducts isolated from the electrochemical reaction of DB[a,l]P and Ade in DMF were identified as DB[a,l]P-10-N1Ade (47%), DB[a,l]P-10-N3Ade (5%), DB[a,l]P-10-N7Ade (2%), and DB[a,l]P-10-N6Ade (6%) (Scheme 1 and Figure 1). In Me2SO, the yields of these adducts were 44, 9, 1, and 3%, respectively. Electrochemical oxidation of DB[a,l]P in the presence of dA generated not only DB[a,l]P-10-N7Ade (3%) and DB[a,l]P-10N6dA (26%), as reported previously (9), but also DB[a,l]P-10N1Ade (25%) (Scheme 2 and Figure 2).


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Figure 2. HPLC separation of the adducts formed by electrochemical oxidation of DB[a,l]P in the presence of dA: (A) CH3OH/H2O gradient and (B) CH3CN/C2H5OH/H2O gradient.

Figure 1. HPLC separation of the adducts formed by electrochemical oxidation of DB[a,l]P in the presence of Ade: (A) CH3OH/H2O gradient and (B) CH3CN/C2H5OH/H2O gradient. (1) DB[a,l]P-10-N1Ade: UV λmax 240, 271, 294, 308, 320, 366, 385, 405, 426 nm; 1H NMR δ 7.37 (d, 1 H, 9-H, J8,9 ) 9.0 Hz), 7.58 (d, 1 H, 11-H, J11,12 ) 7.5 Hz), 7.64 (s, 1 H, 8-H of Ade), 7.81 (t, 1 H, 12-H, J11,12 ) 7.5 Hz, J12,13 ) 7.5 Hz), 7.897.99 (m, 3 H, 2-H, 3-H, 13-H), 8.09 (d, 1 H, 8-H, J8,9 ) 9.0 Hz), 8.21 (t, 1 H, 6-H, J5,6 ) 8.5 Hz, J6,7 ) 7.5 Hz), 8.34 (d, 1 H, 7-H, J6,7 ) 7.5 Hz), 8.37 (bs, 1 H, NH of Ade, can be exchanged with D2O), 8.48 (bs, 1 H, NH of Ade, can be exchanged with D2O), 8.73 (s, 1 H, 2-H of Ade), 9.12 (d, 1 H, 4-H, J3,4 ) 8.5 Hz), 9.23 (d, 1 H, 1-H, J1,2 ) 7.5 Hz), 9.28 (d, 1 H, 5-H, J5,6 ) 8.5 Hz), 9.31 (d, 1 H, 14-H, J13,14 ) 8.5 Hz). (2) DB[a,l]P-10-N3Ade: UV λmax 240, 268, 295, 306, 319, 366, 383, 402, 422 nm; 1H NMR δ 7.29 (d, 1 H, 9-H, J8,9 ) 9.0 Hz), 7.50 (d, 1 H, 11-H, J11,12 ) 8.0 Hz), 7.60 (bs, 2 H, NH2 of Ade, can be exchanged with D2O), 7.82 (t, 1 H, 12-H, J11,12 ) 8.0 Hz, J12,13 ) 6.5 Hz), 7.89-7.98 (m, 3 H, 2-H, 3-H, 13-H), 8.02 (s, 1 H, 8-H of Ade), 8.08 (d, 1 H, 8-H, J8,9 ) 9.0 Hz), 8.20 (t, 1 H, 6-H, J5,6 ) 8.0 Hz, J6,7 ) 7.5 Hz), 8.32 (d, 1 H, 7-H, J6,7 ) 7.5 Hz), 8.64 (s, 1 H, 2-H of Ade), 9.11 (d, 1 H, 4-H, J3,4 ) 8.0 Hz), 9.20 (d, 1 H, 1-H, J1,2 ) 7.5 Hz), 9.26 (d, 1 H, 5-H, J5,6 ) 8.0 Hz), 9.29 (d, 1 H, 14-H, J13,14 ) 9.0 Hz). (3) DB[a,l]P-10-N6Ade: UV λmax 241, 270, 295, 308, 321, 365, 384, 402, 423 nm; 1H NMR δ 6.29 (s, 1 H, 2-H of Ade), 7.76 (t, 1 H, 12-H, J11,12 ) 7.5 Hz), 7.83 (t, 1 H, 13-H, J12,13 ) 8.0 Hz, J13,14 ) 9.0 Hz), 7.86-7.92 (m, 2 H, 2-H, 3-H), 7.98 (d, 1 H, 8-H, J8,9 ) 9.5 Hz), 8.12 (d, 1 H, 9-H, J8,9 ) 9.5 Hz), 8.13 (t, 1 H, 6-H, J5,6 ) 8.5 Hz, J6,7 ) 8.0 Hz), 8.23 (d, 1 H, 7-H, J6,7 ) 8.0 Hz), 8.38 (d, 1 H, 11-H, J11,12 ) 7.5 Hz), 9.08 (d, 1 H, 1-H, J1,2 ) 7.5 Hz), 9.14 (d, 1 H, 4-H, J3,4 ) 10.5 Hz), 9.16 (d, 1 H, 5-H, J5,6 ) 8.5 Hz), 9.21 (d, 1 H, 14-H, J13,14 ) 9.0 Hz). 9.87 (s, 1 H, 8-H of

Ade), 10.38 (bs, 1 H, N6-H of Ade, can be exchanged with D2O), 13.19 (bs, 1 H, 9-H of Ade, can be exchanged with D2O). The exact masses of these adducts were determined by peak matching at high resolving power (R ) 10 000) to be within 1.0 ppm of the theoretical value for C29H18N5. (4) DB[a,l]P-10-N7Ade. According to HPLC retention time, UV, and NMR data, the DB[a,l]P-10-N7Ade obtained by electrochemical oxidation of DB[a,l]P in the presence of dA or Ade was identical to the DB[a,l]P-10-N7Ade obtained previously with dA (10).

Results and Discussion Synthesis of Adducts by Electrochemical Oxidation. Anodic oxidation of DB[a,l]P in the presence of Ade in DMF or Me2SO yielded four adducts: DB[a,l]P-10N1Ade, DB[a,l]P-10-N3Ade, DB[a,l]P-10-N7Ade, and DB[a,l]P-10-N6Ade (Scheme 1). Anodic oxidation of DB[a,l]P in the presence of dA in DMF yielded three adducts: DB[a,l]P-10-N1Ade, DB[a,l]P-10-N7Ade, and DB[a,l]P-10N6dA (Scheme 2). HPLC Purification of Adducts. When the components in the mixture of adducts obtained by oxidation of DB[a,l]P in the presence of Ade were separated by reverse-phase HPLC with the CH3CN/H2O gradient, three sharp peaks, corresponding to DB[a,l]P-10-N3Ade, DB[a,l]P-10-N6Ade, and DB[a,l]P-10-N7Ade, as identified below, were seen (Figure 1A). In addition, a very broad increase in the baseline was observed. When the mixture was assessed by HPLC with a CH3CN/C2H5OH/H2O gradient (Figure 1B), the very broad increase in the baseline became the major peak, and the responsible component was identified as DB[a,l]P-10-N1Ade (see


below). This finding led us to repeat the oxidation of DB[a,l]P in the presence of dA (10), and the DB[a,l]P-10N1Ade was also found to be formed in this reaction (Figure 2B). Under these conditions, however, DB[a,l]P10-N6dA eluted as a very broad peak, and it was necessary to use the CH3CN/H2O gradient to purify DB[a,l]P10-N6dA and DB[a,l]P-10-N7Ade (Figure 2A). Via application of these two gradients, it is possible to separate the four Ade adducts formed by reaction of DB[a,l]P radical cation and Ade and the three adducts formed by reaction of DB[a,l]P radical cation and dA. Use of C2H5OH gradients has allowed us to recover the N1Ade adducts as the major ones formed by several PAHs with Ade (1417). Structure Elucidation of New Adducts. Evidence for the structure of DB[a,l]P-10-N1Ade, DB[a,l]P-10N3Ade, and DB[a,l]P-10-N6Ade was obtained by a combination of UV, NMR, and electrospray MS/MS. Above 300 nm, the UV spectra of the DB[a,l]P-10-N1Ade, DB[a,l]P-10-N3Ade, and DB[a,l]P-10-N6Ade exhibited absorbance maxima red-shifted by 3-13 nm, compared to those for DB[a,l]P; this is characteristic of substitution at C-10 of DB[a,l]P, as was previously observed (10). NMR Spectroscopy. The chemical shifts of the protons of DB[a,l]P-10-N1Ade, DB[a,l]P-10-N3Ade, and DB[a,l]P-10-N6Ade (Figure 3) were assigned by 1H NMR and COSY. The 9-H and 11-H resonances were shifted upfield relative to those of DB[a,l]P (8.12 and 8.42 ppm, respectively, not shown) owing to substitution of Ade at the C-10 position of the parent compound. 1 H NMR of DB[a,l]P-10-N1Ade showed two broad singlets around 8.37 and 8.48 ppm in Me2SO at 25 °C (Figure 3A), which could be exchanged in D2O. Bonding of the C-10 of DB[a,l]P to the N-1 of Ade is suggested by the splitting of the resonances of the NH2 protons. In fact, all of the PAH-N1Ade adducts characterized thus far exhibit this feature (14), and the splitting is attributed to the close proximity of the large aromatic moiety to the NH2 group; this creates a different chemical environment for the two protons owing to a rotational energy barrier. Furthermore, the 2-H of Ade is shifted downfield for the N1Ade adducts linked to an aromatic ring, whereas it is shifted upfield for N1Ade adducts that are linked to a meso-anthracenic methyl group (14). In the NMR spectrum of DB[a,l]P-10-N6dA, one can observe two broad D2O-exchangeable singlets at 10.38 and 13.19 ppm (Figure 3C). The singlet at 10.38 ppm is similar to the NH signal of DB[a,l]P-10-N6dA (10.50 ppm) (10), whereas the singlet at 13.19 ppm is very similar to that of the 9-H in the NMR spectrum of Ade (12.82 ppm, not shown). Because this adduct was obtained by electrochemical oxidation of DB[a,l]P in the presence of Ade, its structure is assigned as DB[a,l]P-10-N6Ade. The structure of DB[a,l]P-10-N3Ade was deduced upon assignment of the 8-H and 2-H proton resonances by NOE analysis of the adduct. In fact, irradiation of the N6 amino group (Figure 3B) leads to a selective NOE enhancement of the 2-H(Ade) signal at 8.64 ppm. Further evidence that the downfield 8.64 ppm proton resonance (8.06 ppm for Ade, not shown) corresponds to that of 2-H(Ade) is the deshielding of the 2-H signal of Ade in DB[a,l]P-10-N1Ade (8.73 ppm); this is similar to the 2-H signal in the BP-6-N3Ade and BP-6-N1Ade adducts [8.60 and 8.68 ppm, respectively (16)]. The structure assignment of DB[a,l]P-10-N3Ade is further corroborated by mass spectrometric studies (see below).


Figure 3. NMR spectra of (A) DB[a,l]P-10-N1Ade, (B) DB[a,l]P-10-N6Ade, and (C) DB[a,l]P-10-N3Ade.

Mass Spectrometry. We recently compared (21) the product ion spectra from MS/MS, using six different approaches, and showed that distinction of the four Ade isomers is most readily achieved via low-energy collisional activation of electrospray-produced ions on a triple quadrupole, or by postsource decompositions of matrixassisted laser desorption ionization-produced ions on a time-of-flight mass spectrometer. Therefore, we present here the spectra of the four isomers introduced by electrospray ionization (ESI), activated by low-energy collisional activation to give products that were analyzed with triple-quadrupole mass spectrometry. These product ion spectra (from the Quattro II instrument) were not published previously, but are nearly identical to those that were (21) and to those recorded on the Finnigan TSQ 7000 triple-quadrupole instrument (see Experimental Methods). The interested reader is referred to ref 21 for the results from the other tandem methods and for additional discussion. All isomers produced, upon collisional activation, the m/z 302 ion as the most abundant fragment (Figure 4). This ion is expected on the basis of previous studies (10), and is the radical cation of the DB[a,l]P species, which is released in a fragmentation process. Adjoining masses of m/z 301 and 300 complete a triplet of fragment ions.


Figure 4. Product ion mass spectra of the ESI-produced [M + H]+ ions of the four isomeric DB[a,l]P adducts with Ade: (A) DB[a,l]P-10-N1Ade, (B) DB[a,l]P-10-N3Ade, (C) DB[a,l]P-10-N6Ade, and (D) DB[a,l]P-10-N7Ade. The spectra were taken with a Micromass Quattro II triple-quadrupole mass spectrometer with a collision energy of 40 eV. The spectra are nearly identical with those taken with a Finnigan TSQ 7000 (see Experimental Methods) and those published earlier (21).

The relative abundances of the members of this triplet are isomerically specific, but it is difficult to understand the chemical basis for the differences. Nevertheless, the pattern can be used as a “fingerprint” once the isomers have been identified. The extent of ammonia loss is also isomerically specific for some of the isomers, and the loss is always more facile for the isomers in which the exocyclic NH2 of the Ade ring adjoins the PAH ring (i.e., DB[a,l]P-10-N1Ade and DB[a,l]P-10-N7Ade). We suggest that the process is anchimerically assisted by participation of the PAH ring. The ammonia loss is also surprisingly detectable for DB[a,l]P-10-N6Ade, suggesting that the PAH group can migrate to other locations of the Ade ring. The decomposition reaction that clearly distinguishes the isomer with PAH substitution of the exocyclic NH2 of Ade is that which produces the m/z 317 ion, which is only significant in the spectrum of DB[a,l]P-10-N6Ade. This ion is likely to be the ArNH2 radical cation (where Ar is the PAH radical), which can only be formed in the N6 isomer. We observed this to be the case in previous studies (12). Conspicuously missing from the spectrum of the N6 isomer are the ions of m/z 327 and 328, which arise from cycloreversion of a purine ring in eliminating ArNC for the other three isomers. Isomeric distinctions for modified nucleobases in which the 1 or 3 position of Ade is substituted are most difficult, not only by mass spectrometry but also by NMR. A set of reactions that occur upon collisional activation, how-

Li et al.

ever, presents a clear-cut distinction. Unsubstituted Ade is known to fragment in tandem mass spectrometry by losing NH3, followed by expulsions of two molecules of HCN (22). As was documented by isotopic labeling, losses of the first and second HCN come mainly from the sixmembered ring of this purine nucleobase, whereas the specificity of the second loss is lower. With substitution at the 1 position, one expects the loss of NH3, but not ready expulsion of two molecules of HCN. This is, in fact, the case for the isomer we call DB[a,l]P-10-N1Ade. The isomer assigned as the 3-substituted one, however, gives loss of ammonia, followed by two expected losses of HCN (to give the ions of m/z 392 and 365). The ion of m/z 340 is also distinctive for the N3 isomer and may indicate a structure in which N-C-N is bonded to the PAH ring and one H atom has been expelled. The losses of NH3 and HCN are expected to be most facile when the PAH is linked to the five-membered ring of Ade, as for DB[a,l]P-10-N7Ade. These reactions also produce distinctive postsource decay spectra of matrix-assisted laser desorption ionization-produced ions, but less so for fast atom bombardment-produced ions that are activated by highenergy collisional activation (21). FLNS. The prowess of FLNS has been demonstrated in many in vitro and in vivo studies of DNA damage from carcinogenic PAH such as BP, 7,12-dimethylbenz[a]anthracene, and DB[a,l]P (20). FLNS is utilized here to provide fingerprint identification spectra for DB[a,l]Pderived Ade adducts. It is capable of distinguishing between a given PAH covalently bound to different DNA bases and to different nucleophilic centers of a given base (20). The 77 K fluorescence spectra obtained under non-linenarrowing conditions (λex ) 308 nm) for the DB[a,l]PAde one-electron oxidation adducts are very similar, with their (0,0) origin bands located at approximately 423 nm (data not shown). Although minor differences were observed [e.g., for DB[a,l]P-10-N1Ade, the (0,0) band is centered at 423.1 nm, DB[a,l]P-10-N3Ade at 422.9 nm, DB[a,l]P-10-N7Ade at 423.6 nm, and DB[a,l]P-10-N6Ade and DB[a,l]P-10-N6dA at 423.2 nm], the resolution is not sufficient for adduct identification. A distinction between the N6Ade and N6dA adducts and the other three adducts is that the former have broader (0,0) fluorescence origin bands (full width at half-maximum bandwidth of 260 cm-1) than the other three (full width at half-maximum bandwidth of 170 cm-1), indicating stronger electronic coupling with the matrix, as discussed below. To distinguish these adducts, one must employ vibronic excitation. Vibronically excited FLN spectra for DB[a,l]P10-N1Ade, DB[a,l]P-10-N3Ade, DB[a,l]P-10-N6Ade, and DB[a,l]P-10-N7Ade, obtained using a series of excitation wavelengths ranging from 418.5 to 404.0 nm, exhibit differences that provide identification (fingerprinting) of these adducts (see Figure 5 for an example with an excitation wavelength of 416.0 nm). By comparison of the FLN vibrational frequency patterns shown in Figure 5, it is clear that all four adducts are easily distinguishable. For a given excitation wavelength, the lower-frequency vibrational modes of the N3Ade adduct and the higherfrequency modes of the N7Ade adduct are pumped more efficiently than the corresponding modes for N1Ade. This is shown in Figure 5 (spectra b, d, and a, respectively) as a difference in the vibronic mode intensity distribution for these three adducts, resulting from the shift in the position of the (0,0) fluorescence origin bands. In addition,


Figure 5. FLN spectra of DB[a,l]P-10-N1Ade (a), DB[a,l]P10-N3Ade (b), DB[a,l]P-10-N6Ade (c), and DB[a,l]P-10-N7Ade (d), in C2H5OH, obtained for an excitation wavelength of 416.0 nm. The zero-phonon lines are labeled with their excited state (S1) vibrational frequencies.

there are vibronic frequency shifts, as well as the presence (or absence) of some modes that enable spectroscopic identification. A mode at 360 cm-1 for N1Ade and at 358 cm-1 for N7Ade is shifted to 353 cm-1 for the N3Ade adduct. The N1Ade adduct has a mode at 404 cm-1, N3Ade at 408 cm-1, and N7Ade at 410 cm-1. There is a mode at 373 cm-1 for N3Ade and at 375 cm-1 for N7Ade, but no corresponding mode for N1Ade. There is a mode at 419 cm-1 for N1Ade and N7Ade, but no corresponding mode for N3Ade. The FLN spectra obtained using other excitation wavelengths (data not shown) also exhibit differences in the frequencies of vibronic modes and their intensity distribution for these three adducts. Spectrum c in Figure 5 shows that the FLN spectrum for the N6Ade adduct is distinctly different from those for the N1Ade, N3Ade, and N7Ade adducts. The intensities of the zero phonon line in spectrum c are relatively weak and superimposed on a strong broad-band emission at the low-energy side of the spectrum. This supports our assignment that the broader (0,0) origin band for N6Ade (in the 77 K fluorescence spectrum) is due to stronger electron-phonon coupling with the glass matrix. The stronger coupling is likely due to a different structural conformation for the N6Ade adduct compared to those of the N1Ade, N3Ade, and N7Ade adducts. The DB[a,l]P chromophore is bound to the exocyclic nitrogen of Ade in the N6Ade adduct, whereas it is bound to a nitrogen atom in the Ade ring for the other three adducts. The resulting broader FLN spectrum for N6Ade and some differences in vibrational frequencies provide a means of distinguishing N6Ade (Figure 5, spectrum c) from the other three adducts (spectra a, b, and d). The FLN spectra of the N6Ade and N6dA adducts are very similar, indicating that the influence of the deoxyribose moiety on the fluorescence characteristics of these two adducts is, as expected, very small. Nevertheless, minor differences were observed in the low-frequency mode region. This is shown in Figure 6, which compares FLN spectra for these two adducts obtained using excita-


Figure 6. FLN spectra of DB[a,l]P-10-N6Ade and DB[a,l]P10-N6dA in C2H5OH, obtained at excitation wavelengths of 415.5 (a and b) and 412.0 nm (c and d). The zero-phonon lines are labeled with their excited state (S1) vibrational frequencies. Table 1. Excited State (S1) Vibrational Frequencies (nm) for DB[a,l]P Adductsa N1Ade

N3Ade

N7Ade

283 300 317

286 301 315 342 353 373 394 408

288 301 317 342 358 375 392 410 419

360 391 404 419 442

445

482 488 521 531 541 557

476 489 523 533

601 612 620 647 a

556 582 603 624 640 653

439 450 474 489 521 534 556 575 598 611 620 632 648 657

N6Ade

N6dA

303

299

345 362

344 360

399

399

414 428 441

428 442

478

480

528 535

530

560

547 556

614 623 631

614 622 631

Vibrational frequencies of weak modes have not been included.

tion wavelengths of 415.5 and 412.0 nm. The N6Ade adduct has a mode at 414 cm-1 that is not observed for N6dA (compare spectra a and b in Figure 6). The N6Ade adduct has a characteristic doublet at 528 and 535 cm-1, whereas N6dA has only a single mode at 530 cm-1; there is also a shift in one mode, 560 cm-1 for N6Ade and 556 cm-1 for N6dA (compare spectra c and d in Figure 6). Of the excited state (S1) vibrational frequencies for the five DB[a,l]P-Ade adducts, obtained from their FLN spectra (Table 1), only modes in the 300-700 cm-1 region have been included, because we established that these low-frequency modes are most useful for characterizing structurally similar adducts. The accuracy of the tabulated frequencies is better than (2 cm-1, so frequency differences of 4-5 cm-1 (or greater) are significant. [Some


differences between these tabulated frequencies and values published previously (6) are due to solvent effects; C2H5OH was used as the glass-forming matrix in the present work, whereas a mixture of glycerol, H2O, and C2H5OH was used previously.] There are similarities in many of the modes for these five adducts, as expected, because the adducts are structurally very similar. There are, however, frequency shifts and the presence or absence of particular modes, discussed above and evident in Table 1, that can be used for unambiguous identification of each of the five adducts. From Table 1 and the spectra presented in Figures 5 and 6, it is clear that FLNS provides a means of distinguishing these adducts. Although each of the individual DB[a,l]P-Ade adducts can be distinguished by FLNS, a mixture of the five adducts cannot be resolved by FLNS alone. We recently developed, however, a new approach for high-resolution detection and characterization of closely related compounds in a chemical separation process, capillary electrophoresis interfaced with FLNS for on-line structural characterization (23, 24), and this will be applied to future studies of isomeric adducts. Relative Reactivity of DB[a,l]P Radical Cation at N-1, N-3, and N-7 of Ade. The reactivity of Ade with DB[a,l]P radical cation depends on two major physicochemical criteria: (1) the nucleophilicity of the N-1, N-3, and N-7 positions and (2) the steric accessibility of these nucleophilic sites to the DB[a,l]P radical cation, which has charge localized at C-10. Electrostatic molecular potential minima at the nucleophilic sites of the four nucleic acid bases have been proposed by Pullman and Pullman as an index of nucleophilicity (25). For the N-1, N-7, and N-3 positions of Ade, the values are -70.4, -67.1, and -62.6 kcal/mol, respectively, suggesting that the nucleophilicity of N-1 is greater than that of N-7, which is greater than that of N-3. When DB[a,l]P radical cation reacts with Ade, however, more adduct is formed at N-3 than at N-7. The smaller reactivity at N-7 is attributed to the lower accessibility of the N-7 position to the DB[a,l]P radical cation, created by the adjacent 6-NH2 group (26). Thus, the relative yields of these three adducts can be rationalized in terms of nucleophilicity and steric accessibility.

Conclusions Electrochemical oxidation of DB[a,l]P in the presence of Ade produces DB[a,l]P-10-N1Ade as the major adduct, and DB[a,l]P-10-N3Ade, DB[a,l]P-10-N6Ade, and DB[a,l]P-10-N7Ade as minor adducts. The ratio of the yields of these adducts reflects the nucleophilic reactivity and steric accessibility of the different nitrogen atoms of Ade in Me2SO or DMF. With dA, the DB[a,l]P radical cation produced only DB[a,l]P-10-N1Ade, DB[a,l]P-10-N7Ade, and DB[a,l]P-10-N6dA. As anticipated, DB[a,l]P-10N3Ade was not obtained, owing to the shielding of the N-3 position of Ade by the deoxyribose ring. Until recently, N1Ade adducts, including DB[a,l]P-10-N1Ade, were not isolated and identified, owing to the unique physicochemical properties of the N1Ade adducts that cause them to elute very broadly with typical HPLC eluents, CH3OH/H2O (Figures 1 and 2) and CH3CN/H2O gradients. The use of a CH3CN/C2H5OH/H2O HPLC gradient (Figures 1 and 2) allowed us to isolate DB[a,l]P10-N1Ade. The N1Ade adduct, as anticipated, has not been found biologically, owing to the unavailability of the

Li et al.

N-1 position in DNA, whereas the DB[a,l]P-10-N3Ade adduct has been found to be abundant in vitro (6) and in vivo.2 Thus far, N3Ade adducts of two other carcinogens, BP (16) and dibenzo[c,g]carbazole (17), have also been reported. The apparent inconsistency of the formation of N3Ade adducts by reaction of PAH radical cations with DNA, but not with dA, can be rationalized by the interference of the deoxyribose at the adjacent N-9 in dA that hinders electrophilic attack at N-3. In DNA, however, the deoxyribose-phosphate backbone rotates the sugar away from the N-3 position, leaving it accessible to reaction with PAH radical cations.

Acknowledgment. This research was supported by U.S. Public Health Service Grants R01-CA49917 and P01-CA49210 from the National Cancer Institute. Core support at the Eppley Institute was provided by National Cancer Institute Grant CP30-CA367271. The Mass Spectrometry Research Resource was supported by the NIH National Center for Research Resources (Grant P41RR00954).

References (1) Cavalieri, E. L., Rogan, E. G., Higginbotham, S., Cremonesi, P., and Salmasi, S. (1989) Tumor-initiating activity in mouse skin and carcinogenicity in rat mammary gland of dibenzo[a]pyrenes: The very potent activity of dibenzo[a,l]pyrene. J. Cancer Res. Clin. Oncol. 115, 67-72. (2) Cavalieri, E. L., Higginbotham, S., RamaKrishna, N. V. S., Devanesan, P. D., Todorovic, R., Rogan, E. G., and Salmasi, S. (1991) Comparative dose-response tumorigenicity studies of dibenzo[a,l]pyrene versus 7,12-dimethylbenz[a]anthracene, benzo[a]pyrene and two dibenzo[a,l]pyrene dihydrodiols in mouse skin and rat mammary gland. Carcinogenesis 12, 1939-1944. (3) Higginbotham, S., RamaKrishna, N. V. S., Johansson, S., Rogan, E. G., and Salmasi, S. (1993) Tumor-initiating activity and carcinogenicity of dibenzo[a,l]pyrene versus 7,12-dimethylbenz[a]anthracene and benzo[a]pyrene at low doses in mouse skin. Carcinogenesis 14, 875-878. (4) LaVoie, E. J., He, J.-M., Meegalla, R. L., and Weyand, E. H. (1993) Exceptional tumor-initiating activitiy of the 4-fluorobenzo[j]fluoranthene on mouse skin: comparison with benzo[j]fluoranthene, 10-fluorobenzo[j]fluoranthene, benzo[a]pyrene, dibenzo[a,l]pyrene and 7,12-dimethylbenz[a]anthracene. Cancer Lett. 70, 7-14. (5) Cavalieri, E., Higginbotham, S., and Rogan, E. (1994) Dibenzo[a,l]pyrene: The most potent carcinogenic aromatic hydrocarbon. Polycyclic Aromat. Compd. 6, 177-183. (6) Li, K.-M., Todorovic, R., Rogan, E. G., Cavalieri, E. L., Ariese, F., Suh, M., Jankowiak, R., and Small, G. J. (1995) Identification and quantitation of dibenzo[a,l]pyrene-DNA adducts formed by rat liver microsomes in vitro: Preponderance of depurinating adducts. Biochemistry 34, 8043-8049. (7) 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. (8) Li, K.-M., George, M., Gross, M. L., Seidel, A., Luch, A., Rogan, E. G., and Cavalieri, E. L. (1999) Structure elucidation of the adducts formed by fjord region dibenzo[a,l]pyrene 11,12-dihydrodiol 13,14-epoxides with deoxyadenosine. Chem. Res. Toxicol. 12, 758-767. (9) Li, K.-M., George, M., Gross, M. L., Lin, C.-H., Jankowiak, R., Small, G. J., Seidel, A., Kroth, H., Rogan, E. G., and Cavalieri, E. L. (1999) Structure elucidation of the adducts formed by fjord region dibenzo[a,l]pyrene 11,12-dihydrodiol 13,14-epoxides with deoxyguanosine. Chem. Res. Toxicol. 12, 778-788. (10) RamaKrishna, N. V. S., Padmavathi, N. S., Cavalieri, E. L., Rogan, E. G., Cerny, R. L., and Gross, M. L. (1993) Synthesis and structure elucidation of the adducts formed by electrochemical oxidation of the potent carcinogen dibenzo[a,l]pyrene in the presence of nucleosides. Chem. Res. Toxicol. 6, 554-560. (11) Rogan, E., Cavalieri, E., Tibbels, S., Cremonesi, P., Warner, C., Nagel, D., Tomer, K., Cerny, R., and Gross, M. (1988) Synthesis and identification of benzo[a]pyrene-guanine nucleoside adducts formed by electrochemical oxidation and horseradish peroxidase-


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catalyzed reaction of benzo[a]pyrene with DNA. J. Am. Chem. Soc. 110, 4023-4029. RamaKrishna, N. V. S., Gao, F., Padmavathi, N. S., Cavalieri, E. L., Rogan, E. G., Cerny, R. L., and Gross, M. L. (1992) Model adducts of benzo[a]pyrene and nucleosides formed from its radical cation and diol epoxide. Chem. Res. Toxicol. 5, 293-302. RamaKrishna, N. V. S., Cavalieri, E. L., Rogan, E. G., Dolnikowski, G. G., Cerny, R. L., Gross, M. L., Jeong, H., Jankowiak, R., and Small, G. J. (1992) Synthesis and structure determination of the adducts of the potent carcinogen 7,12-dimethylbenz[a]anthracene and deoxyribonucleosides formed by electrochemical oxidation: Models for metabolic activation by one-electron oxidation. J. Am. Chem. Soc. 114, 1863-1874. Mulder, P. P. J., Chen, L., Sekhar, B. C., George, M., Gross, M. L., Rogan, E. G., and Cavalieri, E. L. (1996) Synthesis and structure determination of the adducts formed by electrochemical oxidation of 1,2,3,4-tetrahydro-7,12-dimethylbenz[a]anthracene in the presence of deoxyribonucleosides or adenine. Chem. Res. Toxicol. 9, 1264-1277. Hanson, A. A., Rogan, E. G., and Cavalieri, E. L. (1998) Synthesis of adducts formed by iodine oxidation of aromatic hydrocarbons in the presence of deoxyribonucleosides and nucleobases. Chem. Res. Toxicol. 11, 1201-1208. Chen, L., Devanesan, P. D., Higginbotham, S., Ariese, F., Jankowiak, R., Small, G. J., Rogan, E. G., and Cavalieri, E. L. (1996) Expanded analysis of benzo[a]pyrene-DNA adducts formed in vitro and in mouse skin: Their significance in tumor initiation. Chem. Res. Toxicol. 9, 897-903. Chen, L., Devanesan, P. D., Byun, J., Gooden, J. K., Gross, M. L., Rogan, E. G., and Cavalieri, E. L. (1997) Synthesis of depurinating DNA adducts formed by one-electron oxidation of 7H-dibenzo[c,g]carbazole and identification of these adducts after activation with rat liver microsomes. Chem. Res. Toxicol. 10, 225233.

Chem. Res. Toxicol., Vol. 12, No. 9, 1999 757 (18) NIH Guidelines for the Laboratory Use of Chemical Carcinogens (1981) NIH Publication 81-2385, U.S. Government Printing Office, Washington, DC. (19) Gross, M. L., Chess, E. K., Lyon, P. A., Crow, F. W., Evan, S., and Tudge, H. (1982) Triple analyzer mass spectrometry for highresolution MS/MS studies. Int. J. Mass Spectrom. Ion Phys. 42, 243-254. (20) Jankowiak, R., and Small, J. G. (1998) Analysis of PAH-DNA adducts: fluorescence line-narrowing spectroscopy. In The Handbook of Environmental Chemistry, Vol. 3J: PAHs and Related Compounds, (Neilson, A. H., Ed.) pp 119-145, Springer, Heidelberg, Germany. (21) Byun, J., Gooden, J., Ramanathan, R., Li, K.-M., Cavalieri, E. L., 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. 8, 977-986. (22) Nelson, C. C., and McCloskey, J. A. (1992) Collision-induced dissociation of adenine. J. Am. Chem. Soc. 114, 3661-3668. (23) Jankowiak, R., Zamzow, D., Ding, W., and Small, G. J. (1996) Capillary electrophoresis-fluorescence line narrowing (CE-FLNS) system for on-line structural characterization of molecular analytes. Anal. Chem. 68, 2549-2553. (24) Zamzow, D., Lin, L.-H., Small, G. J., and Jankowiak, R. (1997) Application of capillary electrophoresis-fluorescence line narrowing system for on-line spectral characterization of closely related analytes. J. Chromat. A781, 73-80. (25) Pullman, A., and Pullman, B. (1981) Molecular electrostatic potential of nucleic acids. Q. Rev. Biophys. 14, 289-380. (26) 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.

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Synthesis and Structure Determination of the Adducts Formed by

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