UVA-Induced DNA Single-Strand Cleavage by 1-Hydroxypyrene and

1-Hydroxypyrene and Formation of Covalent Adducts ... we have found that, upon UVA irradiation, HOP causes DNA single-strand cleavages and forms...
0 downloads 0 Views 141KB Size
Download PDF
Chem. Res. Toxicol. 2000, 13, 585-593

585

UVA-Induced DNA Single-Strand Cleavage by 1-Hydroxypyrene and Formation of Covalent Adducts between DNA and 1-Hydroxypyrene Shiming Dong,† Huey-Min Hwang,‡ Xiaochun Shi,‡ Laketa Holloway,† and Hongtao Yu*,† Departments of Chemistry and Biology, Jackson State University, Jackson, Mississippi 39217 Received December 2, 1999

1-Hydroxypyrene (HOP), a metabolite found in the urine of humans and laboratory animals exposed to polycyclic aromatic hydrocarbons (PAHs), is known to be both acutely toxic and genotoxic. It has been widely used as a biomarker for studying PAH exposure. In this research, we have found that, upon UVA irradiation, HOP causes DNA single-strand cleavages and forms HOP-DNA covalent adducts. The UVA-induced cleavage of supercoiled plasmid ΦX174 DNA is dependent upon both HOP concentration and UVA dosage. A longer irradiation time or higher HOP concentration induces more DNA cleavage. Results of the photocleavage experiments carried out in the presence of reactive oxygen species scavengers, histidine, sodium azide, mannitol, SOD, and desferal indicate that both the superoxide free radical and singlet oxygen are likely involved in causing DNA single-strand cleavage. The photocleavage is inhibited by the presence of an excited singlet-state quencher, KI, indicating that it is an excited-state reaction. Along with light-induced DNA cleavage, HOP also forms DNA covalent adducts while being degraded upon light irradiation. Light-induced degradation of 20 µM HOP follows firstorder reaction kinetics in a 10% methanolic buffer (10 mM phosphate) solution in the absence or presence of 40 µM calf thymus DNA, with degradation half-lives of 20 or 15 min, respectively. The shorter degradation half-life in the presence of DNA is due to the formation of the HOPDNA covalent adduct. The formation of the HOP-DNA covalent adduct is evidenced by comparing the UV-vis absorption and fluorescence emission spectra of the pure HOP with those of the HOP-DNA adduct. The covalent HOP-DNA adduct produced due to irradiation was purified by either extensive dialysis (3 × 500 mL buffer solutions), phenol and chloroform extraction followed by ethanol precipitation, or chloroform extraction alone. The isolated HOPDNA adduct has an absorption peak at 353 nm, which is 8 nm red-shifted compared to that of free HOP. The fluorescence emission for HOP-DNA is at least 70 times weaker than that for free HOP in solution. In summary, the findings with HOP reveal that, in addition to metabolic activation that eventually leads to the formation of alkylated DNA adducts or other forms of DNA damage, HOP may be activated by light to produce DNA single-strand cleavage and covalent DNA adducts. These DNA lesions can be sources of toxicity.

Introduction (PAHs)1

Polycyclic aromatic hydrocarbons are a class of compounds produced from forest fire, volcanic eruption, tobacco smoke, food processing, and incomplete burning of fuel and other materials (1, 2). Being ubiquitous in the environment, PAHs are also thought to induce cancer tumors, primarily in the lungs, bladder, and skin (2-8). The International Agency for Research on Cancer and the United States Environmental Protection Agency have classified some of these compounds as probable human carcinogens (9, 10). * To whom correspondence should be addressed: Department of Chemistry, Jackson State University, Jackson, MS 39217. Telephone: (601) 979-3727. Fax: (601) 979-3674. E-mail: [email protected]. † Department of Chemistry. ‡ Department of Biology. 1 Abbreviations: HOP, 1-hydroxypyrene; HOP-DNA, covalent adduct with DNA and HOP (exact structure not yet determined); UVA, ultraviolet light in the range of 320-400 nm; PAH, polycyclic aromatic hydrocarbon; ct-DNA, calf thymus DNA; sc-DNA, supercoiled form I DNA; oc-DNA, open circular (relaxed) form II DNA; SOD, superoxide dismutase; TBE, Tris-borate EDTA buffer; BPDE, benzo[a]pyrene diol epoxide; ROS, reactive oxygen species.

PAHs themselves are relatively inert molecules and are generally considered nontoxic. It is their transformation products that are active and have been shown to be both acutely and chronically toxic (1, 3, 7, 11-15). The main transformation pathway that turns the relatively inert PAHs into reactive intermediates is metabolism. After entering the cell, PAHs are metabolized into diol epoxides, diones, and other reactive intermediates that react with cellular DNA to form PAH-DNA covalent adducts or cause other forms of DNA damage (2-5, 12, 13, 15-18). The structures of some of the covalent adducts have been well-studied (14). The PAH-DNA covalent adduct formation is considered a source of carcinogenicity and toxicity (1, 3, 7, 12, 13). Oxidative damage to DNA generated during metabolic activation is also thought to be a source of carcinogenicity and toxicity (11, 18-20). Another pathway through which PAH toxicity is enhanced is light activation. There is evidence that PAH mixtures and individual PAHs exhibit photoinduced toxicity in the natural environment toward microorgan-

10.1021/tx990199x CCC: $19.00 © 2000 American Chemical Society Published on Web 06/17/2000

586

Chem. Res. Toxicol., Vol. 13, No. 7, 2000 Chart 1. Structure of 1-Hydroxypyrene

isms, plants, and other organisms (21-26). It was found in these studies that PAHs are generally more toxic when they are exposed to the simulated solar radiation than if they are kept in the dark. The increase in toxicity due to simulated solar radiation can exceed 100-fold (26). Reports on a well-studied PAH, benzo[a]pyrene, showed that it can form a DNA covalent adduct or cause DNA strand breakage under light (27-34). The presence of benzo[a]pyrene can increase the level of formation of 8-hydroxy-2′-deoxyguanine induced by UV light (35), a compound generated due to oxidative damage to DNA. DNA-bonded benzo[a]pyrene diol epoxide (BPDE) can also cause DNA strand cleavage under intensive laser light (36). It has been suggested that all the types of DNA damage listed above relate to tumor induction and other adverse effects (27, 28, 37, 38). Since the toxicity and carcinogenicity of PAHs are the result of their transformation into reactive intermediates, much work has focused on their metabolic or degraded products. One of the metabolic products, 1-hydroxypyrene (HOP), has been found in the urine of human and laboratory animals exposed to PAHs from a variety of environmental sources (39, 40), including polluted urban air (41), coal tar (42), and a graphite-electrode producing plant (43). Because of its consistent presence in the urine, HOP is widely used as a biomarker to study human PAH exposure (8, 39-41, 43-46). Recently, it has been found that HOP is more toxic than its parent compound pyrene (47) and is both acutely toxic and genotoxic (47, 48). It is also found to inhibit the progesterone receptor-mediated transactivation in yeast (49). Although there is no assessment of the amount of HOP present in the environment, it has been detected in plant leaves (50). Release of HOP into the environment through urine is one contamination source, but other sources might exist. Since HOP is a metabolic product of PAHs, it could be present in human and animal cells and possibly in the bloodstream. Therefore, human exposure to HOP is eminent (40). Previously, we have found that 12 PAHs, including pyrene, chrysene, benzo[a]pyrene, and their substituted derivatives, can cause UVA-induced DNA single-strand cleavage (51). In this research, we report a study of the photochemical reaction of HOP with DNA that produces HOP-DNA covalent adducts and causes DNA single-strand cleavage.

Materials and Methods Caution: 1-Hydroxypyrene is both acutely toxic and genotoxic (47-49) and should be handled according to the NIH Guidelines for Chemical Carcinogens. Reagents, Chemicals, and Instrumentation. HOP was purchased from Aldrich and used without further purification.

Dong et al. A HOP stock solution (1 mM) was prepared in methanol and stored in the refrigerator in the dark. The stock solution was diluted with solvents needed to make any working solutions. ΦX-174 phage DNA (80-90% supercoiled RF-1 or sc-DNA) with a molecular mass of 3.6 × 106 Da and 5386 bp, ct-DNA, desferal, L-histidine, mannitol, ethidium bromide, bromophenol blue, xylene cyanol, and SOD were purchased from Sigma-Aldrich. Agarose, sodium azide, mono- and dibasic sodium phosphate, TRIS base, boric acid, and EDTA were purchased from Fisher Scientific. All solvents that were used were spectroscopic grade. The water that was used (18 Ω) was deionized by a Barnstead Nanopure Infinity water deionization system (Dubuque, IA). UV-vis spectra were recorded on a CARY 300E UV-vis spectrophotometer (Varian). Liquid chromatography was run on a Hewlett-Packard 1100 series HPLC system with diode array and fluorescence detectors. The HPLC column was a Hypersil ODS reverse phase C18 column (4 mm × 250 mm). Fluorescence emission spectra were recorded on a Fluoromax-2 spectrofluorometer (Instruments S. A., Inc.). Unless stated otherwise, the buffer used for all experiments was 10% methanol [10 mM PBS (pH 7.1)]. Methanol (10%) was used for better HOP solubility. Spectrophotometric Titration of HOP by ct-DNA. A solution of 20 µM HOP in the buffer was titrated with sequential aliquots of a 1.33 mM ct-DNA solution containing 20 µM HOP. During the titration, the concentration of HOP was kept constant. After each addition of DNA, the UV-vis absorption spectra were recorded to monitor the interaction. Photodegradation of HOP in the Presence or Absence of DNA. Two sets of 2 mL solutions of 20 µM HOP in the buffer in the presence or absence of 40 µM ct-DNA were filled in two identical quartz cuvettes (10 mm light path). The cuvettes were placed on top of a Pyrex glass support and were irradiated by UVA light from below the Pyrex glass. The lamp was set beneath the Pyrex glass support, and a stream of cold air was blowing toward the bottom of the support during the irradiation period to eliminate any heat generated by light. The light source was a 100 W UV lamp (type B, UVP Inc., Upland, CA) that produces a main emission band with a wavelength of 365 nm. The distance from the surface of the lamp to the Pyrex glass support was 6.5 cm. The intensity of the light output at this distance was determined to be 170 J/cm2‚h using a model PMA 2100 UVA radiometer (Solar Light Co., Inc., Philadelphia, PA). After each irradiation period, an absorption spectrum was recorded on the CARY 300 UV-vis spectrophotometer. Absorption values at 345 nm were used for kinetic analysis. The firstorder kinetic process was analyzed with the equation ln Ao/At ) kt, where Ao and At are the absorption intensity of HOP at 345 nm at time zero and at time t, respectively, and k is the first-order reaction rate constant. A linear plot was obtained for ln Ao/At versus t for the initial 30 min of irradiation. The slope of the plot was the rate constant k, and the half-life (t1/2) for degradation was calculated with the equation t1/2 ) 0.693/k. In the presence of DNA, one degradation pathway for HOP is formation of a DNA covalent adduct (see the later discussion). The HOP covalently linked to DNA also absorbs at 345 nm. Therefore, the absorbance value for HOP-DNA was subtracted for calculating the absorbance value of the nondegraded HOP. UVA-Induced HOP-DNA Covalent Adduct Formation. A 30 mL solution of 20 µM HOP in the buffer in the presence of 100 µM ct-DNA was filled in a 50 mL Kimax beaker. The beaker was placed on top of the Pyrex glass support, and the lamp was set up as described for the previous experiment. Both the Pyrex glass support and the Kimax beaker efficiently filtered off light below 300 nm, but allowed most of the light with longer wavelengths to pass through. This fact was tested via the UV absorption spectra of the glasses. A 2 mL sample was taken at various irradiation time intervals, and all of these samples were divided into two 1 mL portions. One portion was kept in the dark, and the other was filled into individual dialysis tubes and dialyzed against 3 × 500 mL of buffer for a total of 36 h. The dialysis tube was a Spectra/Por membrane with a molecular

PAH Light-Induced DNA Damage and Phototoxicity weight cutoff of 12 000. The absorption spectra of the dialyzed samples were recorded and compared with those of the nondialyzed samples. To confirm that the above dialysis method efficiently removed all noncovalently bound HOP and its photoproducts, the following experiments were conducted. (1) For extraction by chloroform, a 2 mL irradiated sample was extracted by chloroform (3 × 5 mL). The organic and aqueous mixture was stirred vigorously for 20 min for efficient mixing. (2) For phenol and chloroform extraction in combination with ethanol precipitation, a 2 mL irradiated sample was extracted with a 1:24 mixture of phenol and chloroform (3 × 5 mL). The remaining aqueous solution precipitated in cold 60% ethanol. After each treatment, the absorption spectrum for the isolated HOP-DNA adduct was recorded. Detection of the HOP-DNA Adduct by HPLC Coupled with a Diode Array Detector. A 2′-deoxyribonucleotide octamer duplex, d(5′-GCTAGGGC-3′)‚d(5′-GCCCTAGC-3′), was synthesized by Gemini Biotech (The Woodlands, TX). Its Tm is 22 °C in the 10 mM sodium phosphate buffer in 10% methanol (pH 7.1). A 200 µL solution containing 25 µM HOP and 200 µM octamer (in base pairs) in the buffer was placed in a cuvette, and the solution was cooled in an ice-water bath to 0-5 °C in a 100 mL Kimax beaker. At this temperature, the octamer should be in duplex form. The solution was irradiated by light for 4 h using the same setup for the UV lamp described above. After the irradiation, the sample was directly analyzed by HPLC [linear gradient lasting 120 min from 0 to 60% methanol; flow rate of 1 mL/min; the eluent was buffered by 20 mM phosphate (pH 7)]. UVA-Induced Plasmid DNA Cleavage by HOP. Solutions (a total of 60 µL for each sample) containing ΦX174 phage DNA (27 µM), various amounts of HOP, and other added chemicals were added to the wells of a 24-well (3 × 8) flat-bottomed Titertek plate (ICN Biochemicals). The Titertek plate was tightly covered with a piece of glass and placed on the Pyrex glass support as described earlier. The Pyrex glass here also served as a light filter to cut off any light below 300 nm that can potentially damage DNA. The lamp was set the same way as described above for the degradation experiments. The samples were irradiated for 1 h. During the irradiation, the Titertek plate was turned around horizontally every 15 min to eliminate light heterogeneity. Gel Electrophoresis. After irradiation, each well of the Titertek plate was added with 10 µL of a gel-loading dye solution (bromophenol blue and xylene cyanol in 50% glycerol). Then, 10 µL of the sample was loaded into the wells of a pre-prepared 1% agarose gel. The gel was run in 1× TBE buffer (pH 8.27) at 105 V for 70-90 min at ambient temperature. Following electrophoresis, the gel was stained with ethidium bromide (2 mg/L) for 30 min and analyzed using a NucleoVision 740 GelDocumentation System (NucleoTech Inc.). The sc-DNA and the oc-DNA were well separated on the gel with the faster-moving band being the sc-DNA and the slower-moving band being the oc-DNA. There was no sign for the linear DNA form because it would move slower than the sc-DNA but faster than the oc-DNA. The amounts of the sc-DNA and the oc-DNA were quantified by the integrated fluorescence intensities of the bands after subtracting a common background. A coefficient of 1.66 was used to correct the lower efficiency for ethidium bromide binding to the sc-DNA than to the oc-DNA (52). The percentage of sc-DNA, Xsc, was calculated by

Xsc ) Asc/(Asc + Aoc/1.66) × 100% where Asc and Aoc are the integrated total fluorescence intensities for the sc-DNA and oc-DNA bands, respectively. Therefore, the percent of oc-DNA equals 100 - Xsc.

Results Noncovalent Interaction between HOP and DNA. The change in UV-vis spectra for HOP due to the

Chem. Res. Toxicol., Vol. 13, No. 7, 2000 587

Figure 1. Spectrophotometric titration of 20 µM HOP by ctDNA in 10 mM PBS buffer in 10% methanol (pH 7.1). The [ctDNA]/[HOP] ratios were 0, 0.2, 0.4, 0.5, 0.8, 1, 1.5, 2, 3, 5, 7, and 10 (from top to bottom). The concentration of DNA, [ct-DNA], is expressed in base pairs.

addition of DNA is shown in Figure 1. HOP has three main absorption bands at 240, 278, and 345 nm as shown in Figure 2a. The 345 nm band has a shoulder on each side of the peak. There is also a weaker absorption band at 380 nm. Since both the 240 and 278 nm bands overlap strongly with the absorption bands of DNA bases near 260 nm, they were not used for DNA interaction studies. When ct-DNA was titrated into the HOP solution, the magnitudes of the 345 nm absorption band and its left shoulder decreased while the magnitude of its right shoulder increased (Figure 1). The magnitude of the 380 nm band also decreased, and the band shifted slightly to the red. As a result of the titration, two well-defined isobestic points at 353 and 386 nm emerged. There was a third seemingly isobestic point at 379 nm, but it was not as well-defined as the other two isobestic points. These well-defined isobestic points indicate that a noncovalent complex between DNA and HOP is formed under these conditions. However, we were not able to obtain an accurate binding constant because the titration end point could not be reached. To obtain a binding constant using the McGhee and von Hippel analysis (53), one would need the final absorption value, which is the absorption value for the complex, to calculate the binding ratio. Unfortunately, this absorption value for the complex could not be obtained because the titration end point could not be reached. However, if absorption values that are slightly lower than the last absorption value obtained through the titration are assumed, the estimated binding constant for HOP binding to ct-DNA is in the range of 10-100 M-1 (data not shown), well below the values of 104-107 M-1 for many other DNA-interacting agents (54, 55), including benzo[a]pyrene derivatives (56). UVA-Induced Degradation of HOP in the Presence or Absence of ct-DNA. Light-induced degradation of HOP upon UVA irradiation was followed by the spectral change using the 345 nm absorption band. Figure 2 shows the absorption spectra of 20 µM HOP in the solution after it was irradiated for a certain period

588

Chem. Res. Toxicol., Vol. 13, No. 7, 2000

Dong et al.

Figure 3. Light-induced formation of the HOP-DNA covalent adduct. Line 1 represents the spectrum for a 20 µM HOP solution in the presence of 100 µM ct-DNA in buffer [10 mM PBS (pH 7.1) in 10% methanol]. Line 1′ is the spectrum for the solution represented by line 1, but after dialysis for 36 h against the 10% methanolic buffer (3 × 500 mL). Line 2 is the same as line 1, but after irradiation for 60 min under UVA (170 J cm-2 h-1). Line 2′ is the same as line 2, but after dialysis for 36 h against the 10% methanolic buffer (3 × 500 mL).

Figure 2. Photodegradation of HOP in an aerated aqueous solution in the absence (a) or presence (b) of ct-DNA. (a) A 20 µM HOP solution of 10 mM PBS buffer (pH 7.1) in 10% methanol was irradiated by UVA (170 J cm-2 h-1). (b) Same conditions as described for panel a, but in the presence of 40 µM ct-DNA. The irradiation times for both panels a and b were 0, 1.5, 3, 4.5, 6, 9, 12, 15, 21, and 30 min (from top to bottom).

of time in the absence (panel a) or presence (panel b) of 40 µM ct-DNA. It can clearly be seen that the magnitudes of the 240, 278, and 345 nm absorption bands gradually diminish due to light irradiation, indicating that HOP is degraded upon light irradiation. At the same time, a broad but relatively weak band appears between 400 and 500 nm. The appearance of this new band indicates that at least one new photoproduct for HOP is formed due to degradation. The plot of ln Ao/At versus irradiation time, t, resulted in a straight line for the initial 30 min of the irradiation period (data not shown). From the slope of the plot, the first-order rate constant (k) was obtained. Thus, the half-life t1/2 was calculated to be 20 min. In the same way, a t1/2 of 27 min was obtained for the

degradation of 20 µM HOP in the presence of 40 µM ctDNA. If the absorbance for pure HOP were corrected by subtracting the absorbance value at 345 nm for the HOP-DNA covalent adduct produced during the irradiation, a t1/2 of 15 min was obtained. UVA-Induced Formation of the HOP-DNA Covalent Adduct. If the mixture of HOP and DNA were irradiated by UVA light, some HOP molecules would form covalent bonds with DNA. The HOP-DNA covalent adduct is isolated and identified by UV-vis absorption and fluorescence emission spectroscopy. Figure 3 shows the UV-vis absorption spectra of HOP in the presence of DNA before (line 1) and after 60 min (line 2) of irradiation. The lower absorption value for the irradiated sample (line 2) is due to HOP degradation. Lines 1′ and 2′ represent the same samples as the samples for lines 1 and 2, respectively, but after dialysis for 36 h against the 10% methanolic buffer (3 × 500 mL). While all the HOP absorption represented by line 1′ disappeared, an absorption band, which is different from that of free HOP, remained as shown by line 2′. This means that while noncovalently bound HOP molecules are free to diffuse out of the membrane, some of the HOP or its photoproducts must have formed covalent bonds with DNA molecules, which are too large to diffuse out of the membrane. To confirm this result, extraction methods are used in combination with ethanol precipitation to remove any potential HOP or its photoproducts that are not covalently bound to the ct-DNA. These extraction methods have been used to successfully remove molecules such as BPDE (58), nitropyrene and aminopyrene (59), daunorubicin (60), and ethidium (61) that are not covalently bound to DNA from their covalent bound counterparts. The HOP-DNA samples being studied were (1) extracted by chloroform alone or (2) extracted by a mixture of

PAH Light-Induced DNA Damage and Phototoxicity

Figure 4. Fluorescence emission spectra of HOP and the HOP-DNA adduct in the 10% methanolic buffer [10 mM PBS (pH 7.1)]. The excitation wavelength was 353 nm. The spectrum for the HOP-DNA adduct is amplified 50 times because it is too weak to be seen.

phenol and chloroform (1:24) followed by ethanol precipitation. The absorption band for the HOP-DNA adduct at 353 nm persisted after these treatments. Although the intensity of the 353 nm band is not the same for each method, the ratio between the absorption of DNA at 260 nm (A260) and the absorption of the HOPDNA adduct at 353 nm (A363), A260/A353, remained the same no matter which method was used (10 ( 1). This evidence strongly suggests that HOP formed a covalent bond with DNA upon light irradiation. Absorption and Fluorescence Emission Spectra of the HOP-DNA Covalent Adduct. In comparison to that for free HOP in solution in the presence or absence of DNA, the absorption for the HOP-DNA adduct is 8 nm red-shifted as shown in Figure 3. While the absorption maximum for noncovalently bound HOP is 345 nm (line 1), the absorption maximum for the HOP-DNA adduct is at 353 nm (line 2′). The similar red-shifted spectrum was also observed for BPDE upon DNA covalent adduct formation (57). In this case, the absorption maximum at 344 nm for the free BPDE shifts to 354 nm upon formation of a covalent bond with a guanine residue in the duplex DNA. Also, the 353 nm absorption band for the HOP-DNA adduct (Figure 3, line 2′) has some similar features as the 345 nm band for free HOP (Figure 3, line 1) such as the left and right shoulders. The fluorescence emission spectra for both HOP and the HOP-DNA adduct were recorded with an excitation wavelength of 353 nm. HOP has a strong fluorescence emission peak at 384 nm and two weaker emission peaks at 406 and 428 nm (Figure 4). To obtain a fluorescence spectrum for the HOP-DNA adduct, a solution of 5 mL of the HOP-DNA adduct obtained from the degradation experiments described above was dialyzed for a total of 84 h against 6 × 500 mL of buffer in an effort to eliminate any free HOP that is present. The fluorescence emission spectrum for the HOP-DNA adduct in the remaining solution was recorded and plotted on the same graph after a 50-fold enlargement with the emission spectrum for the free HOP (Figure 4). Even after the enlargement,

Chem. Res. Toxicol., Vol. 13, No. 7, 2000 589

the fluorescence intensity for the HOP-DNA adduct is still less than half of that for the free HOP. The area underneath the emission spectrum for the free HOP is 70 times larger than the area for the HOP-DNA adduct before enlargement. Since the absorption intensities for both solutions were made the same for the purpose of this comparison, it is safe to say that the fluorescence quantum yield for the HOP-DNA adduct is at least 70 times less than that of the free HOP under the same experimental conditions. To say the least, it is because it is not absolutely certain that all of the free HOP molecules were completely eliminated through the extensive dialysis. A trace amount of free HOP may dominate the total fluorescence. Therefore, the shape of the emission spectrum for the pure HOP-DNA adduct may be slightly different from what is shown here because of the possible interference by traces of free HOP. Nonetheless, it is clear that the formation of the HOPDNA adduct greatly quenched the fluorescence of HOP. The same fluorescence quenching effect was seen for BPDE upon covalent adduct formation with guanine either as free guanine or in duplex DNA (64). BPDE is strongly fluorescent, but its fluorescence is quenched upon formation of a covalent bond with guanine (64). This observation is also consistent with the conclusion reached above that HOP formed a covalent bond with DNA. Chromatographic Detection of the HOP-DNA Covalent Adduct. If an octamer DNA, d(5′-GCTAGGGC3′)‚d(5′-GCCCTAGC-3′), were irradiated in the presence of HOP, covalent adducts of DNA with HOP would be formed. The HOP-DNA covalent adducts were detected by HPLC since they had retention times different from those of the original DNA strands. At room temperature and under chromatographic conditions, the pure octamer existed as two single strands in the mobile phase shown by the two peaks at 29.5 and 32.8 min on HPLC (data not shown). Via comparison of the chromatograms recorded for each individual strand, it was found that the 29.5 min peak was due to d(GCTAGGGC) and the 32.8 min peak was due to d(GCCCTAGC). If the mixture of the octamer (200 µM) and HOP (20 µM) were irradiated at 0-5 °C for 4 h, new peaks at 38.5, 40.4, and 48.5 min would be detected on HPLC (Figure 5), in addition to some minor peaks. The absorption spectra of these three peaks recorded by the diode array detector are identical and are shown in the inset of Figure 5. This absorption spectrum included contributions from both DNA (absorption near 260 nm) and HOP (or its derivative near 350 nm). The absorption band near 350 nm is nearly identical with the band for the HOP-DNA adduct shown in Figure 3 (line 2′). This strongly indicated that HOP molecules formed a covalent bond with DNA upon light irradiation. There are at least three different HOP-DNA adducts represented by the three peaks in the chromatogram. Structural analysis and characterization of these DNA adducts are underway. Single-Strand DNA Cleavage Is Dependent on both UVA Dosage and HOP Concentration. Supercoiled ΦX174 DNA (27 µM in base pairs), mixed with 0.5 µM HOP, was irradiated by UVA light, and a series of samples from the mixture were taken at various time intervals and run on a 1% agarose gel. Figure 6 is a photograph of the gel. Above the gel is the quantified band intensity for oc-DNA. As the irradiation time becomes longer, the band intensity for oc-DNA increases. At the same time, the band intensity for sc-DNA de-

590

Chem. Res. Toxicol., Vol. 13, No. 7, 2000

Figure 5. HPLC chromatogram of a mixture of a DNA octamer, d(5′-GCTAGGGC-3′)‚d(5′-GCCCTAGC-3′), and HOP after light irradiation. A solution of 200 µM octamer (in base pairs) and 25 µM HOP in the 10% methanolic buffer [20 mM phosphate (pH 7.1)] was irradiated for 4 h in an ice-watercooled bath using the UVA lamp. The HPLC column was a Hypersil ODS C18 column (4 mm × 250 mm, 5 µm). The flow rate was 1 mL/min with a linear gradient of 0 to 60% methanol buffered with 20 mM phosphate at pH 7.1. The inset shows the absorption spectrum for the peak at 48.5 min captured by a diode array detector.

Figure 6. Time course of photoinduced DNA cleavage by HOP in an aerated aqueous solution. A solution containing ΦX174 phage DNA (27 µM) and HOP (0.5 µM) was irradiated with the UVA lamp (170 J cm-2 h-1) for 0, 5, 10, 20, 30, 45, 60, and 90 min (corresponding to lanes 1-8). The solvent was a 10 mM PBS buffer (pH 7.1) with 10% methanol. After irradiation, the sample was loaded onto a 1% agarose gel and run in 1× TBE buffer for 100 min. Then the gel was stained for 30 min with ethidium bromide (2 mg/L), visualized, and quantified with a NucleoVision Gel-Documentation System. The bar graph represents the percentage of oc-DNA in each lane.

creases, indicating that sc-DNA is converted to oc-DNA due to irradiation. The longer the irradiation time, the more sc-DNA is converted to oc-DNA. This sc-DNA to ocDNA conversion must be due to a single-strand DNA cleavage, not a double-strand DNA cleavage. It is known that sc-DNA converts to oc-DNA upon a single-strand cleavage (or nick) and to linear DNA upon a doublestrand cleavage. There is no sign in the gel of the formation of a linear DNA.

Dong et al.

Figure 7. Concentration dependence of UVA-induced DNA cleavage by HOP in an aerated aqueous solution. The 27 µM ΦX174 phage DNA in the presence of various amounts of HOP was irradiated with the UVA lamp (170 J cm-2 h-1) in a 10 mM PBS buffer (pH 7.1) with 10% methanol. After irradiation, the sample was loaded onto a 1% agarose gel and run in 1× TBE buffer for 100 min. Then the gel was stained for 30 min with ethidium bromide (2 mg/L), visualized, and quantified with a NucleoVision Gel-Documentation System. Lanes 1-3 are controls: lane 1, only DNA present and no irradiation; lane 2, DNA and 4 µM HOP kept in the dark; and lane 3, only DNA present and with irradiation for 1 h. Lanes 4-9 were irradiated for 1 h in the presence of increasing amounts of HOP: 0.1, 0.2, 0.4, 0.6, 1, and 2 µM, respectively. The bar graph represents the percentage of oc-DNA in each lane.

The light-induced DNA cleavage is dependent of HOP concentration as well. This was tested by irradiating the mixtures of DNA with various amounts of HOP for 1 h and analyzed by agarose gel electrophoresis. Figure 7 is the photograph of the gel for DNA treated with HOP and UVA. The bands representing the two forms of DNA were quantified, and the percent of oc-DNA was shown as a bar graph above the gel (Figure 7). As the concentration of HOP increased, the amount of sc-DNA decreased, but the amount of oc-DNA increased due to single-strand cleavage caused by HOP and UVA light. Control experiments showed that both UVA light and HOP are necessary for DNA cleavage. In comparison to the amount of original DNA (lane 1), the relative amount of sc-DNA and oc-DNA was not changed regardless of whether the DNA was irradiated by UVA light in the absence of HOP (lane 3) or the DNA was kept in the dark in the presence of HOP (lane 2). Effect of Oxygen on DNA Photocleavage Induced by HOP. To know whether any reactive oxygen species (ROS) is involved in causing DNA cleavage, the DNA cleavage experiments were carried out in the presence of ROS scavengers. Table 1 lists the effects of those scavengers on DNA cleavage. All experiments were carried out with 27 µM DNA and 0.5 µM HOP, and the mixture was irradiated for 1 h. The experiments carried out under argon produced 46% less oc-DNA than in the aerated solution. This confirms that at least one of the reactive oxygen species is involved in causing DNA cleavage. In fact, it seems that both superoxide free radical (O2•-) and singlet oxygen (1O2) are involved. The presence of 50 mM NaN3, a singlet oxygen scavenger, inhibits 43% of the DNA cleavage. If the experiments

PAH Light-Induced DNA Damage and Phototoxicity Table 1. Effect of Scavengers on DNA Photocleavage by HOPa scavenger

ROS quenched

effect on DNA photocleavage (%)

argon histidine, 50 mM mannitol, 50 mM desferal, 10 mM SOD, 200 units/mL NaN3, 50 mM D2O, 100% KI, 50 mM

O2 OH•/1O2 OH• Fe3+ (H2O2) O2•1O 2 enhanced 1O2 lifetime excited singlet state

-(46 ( 2) no effect no effect no effect -(37 ( 3) -(43 ( 3) 63 ( 4 -(67 ( 7)

a Sample solutions containing ΦX174 phage DNA (27 µM in base pairs) and HOP (0.5 µM) were added with various scavengers. The solutions were irradiated for 1 h by UVA at 170 J/cm2 in 10% methanolic buffer [10 mM PBS (pH 7.1)]. Triplicate experiments were carried out to obtain the final average numbers. The experiments under argon were carried out in a septum-sealed cuvette degassed with argon for 15 min before irradiation. Several concentrations of desferal were used to study its effect on DNA cleavage, but no significant effect was detected. The D2O experiments were conducted in buffer prepared by D2O and CD3OD and nondeuterated phosphates.

were carried out in D2O instead of in H2O, the photocleavage was enhanced by 63%. It is known that singlet oxygen has a 10-fold longer lifetime in D2O than in H2O (62). Both of these observations point to the presence of singlet oxygen, although it is still not clear how singlet oxygen causes DNA cleavage. The presence of 200 units/ mL SOD in the experimental mixture inhibits 37% of the DNA cleavage, indicating that O2•- is also likely involved since SOD would convert the reactive O2•- into less reactive species. Hydroxyl free radical and hydrogen peroxide are not likely involved in the DNA cleavage because the cleavage is not affected by the presence of OH• or H2O2 scavengers such as histidine, desferal, and mannitol. Among them, desferal is an agent used to chelate traces of ferric ions that might be present in the solution. A trace amount of ferric ions was detected in other systems and was thought to catalyze the Fenton reaction that converts hydrogen peroxide into the reactive hydroxyl free radicals. Those hydroxyl free radicals in turn cleave DNA (63). Effect of an Excited Singlet-State Quencher on DNA Photocleavage. Iodide ion is an excited singletstate quencher that works by enhancing the intersystem crossing rate from an excited singlet state to the triplet state of a chromophore (65). Indeed, KI quenches the fluorescence of HOP because addition of KI to the solution of HOP reduces the fluorescence intensity (data not shown). The presence of 50 mM KI quenches the photoinduced DNA cleavage by 67%, indicating that the singlet excited state of HOP is involved in causing DNA cleavage. It is known that KI enhances the intersystem crossing from the excited singlet state to the triplet state (65). As a result, the population of the excited singlet state decreases and that of the excited triplet state increases. If the DNA cleavage reaction were related to the tripletstate, it would have been enhanced in the presence of iodide ions. The fact that the presence of iodide ions diminishes the extent of DNA cleavage indicates that it is an excited singlet-state reaction.

Discussion As the results indicated, HOP forms covalent adducts with DNA and causes DNA single-strand cleavage upon UVA irradiation. Photochemically, HOP itself degrades

Chem. Res. Toxicol., Vol. 13, No. 7, 2000 591

quickly in a 10% methanolic buffer with a half-life of 20 min. In the presence of ct-DNA, though, the photodegradation of HOP is accompanied by HOP-DNA covalent adduct formation. As a result, the overall degradation rate for HOP in the presence of DNA increased as shown by a shorter half-life of 15 min. The covalent HOP-DNA adduct, formed by irradiation of a DNA and HOP mixture, was positively identified by the absorption spectrum of the HOP-DNA adduct isolated through dialysis or extraction followed by ethanol precipitation of the adduct formed with ct-DNA, and chromatographic analysis of the adduct formed with a DNA octamer. The 353 nm absorption band for the HOP-DNA covalent adduct could not be removed from DNA by either dialysis or extraction followed by ethanol precipitation. These extraction methods have been utilized to efficiently remove noncovalently bound molecules such as BPDE (58), nitropyrene and aminopyrene (59), daunorubicin (60), and ethidium (61) from large DNA molecules. Also, at least three adducts were identified for the UVAirradiated sample containing a DNA octamer, d(5′GCTAGGGC-3′)‚d(5′-GCCCTAGC-3′), and HOP. Therefore, all the evidence confirms that HOP forms covalent adducts with DNA upon light irradiation. The nature of the covalent bond for the HOP-DNA adduct is not known at this point, and it requires further in-depth studies using methods such as LC/MS and NMR if a sufficient sample can be isolated. However, we may compare it with other known DNA covalent adducts. PAHs have been known to be covalently attached to DNA bases through two enzymatic pathways: the formation of diol epoxides (3, 13, 14) or one-electron oxidation to free radicals (12, 16, 18-20, 22). The metabolic diol epoxides usually form a covalent bond to guanine’s N7 (14), while the one-electron oxidation products, the cation radicals, usually form a covalent attachment to guanine or adenine at different ring positions (12, 19). In addition, there have been observations that benzo[a]pyrene can form a covalent bond with thymine upon light or chemical activation (30, 33). In analogy to these examples, UVA light-activated HOP molecules possibly form covalent bonds to DNA bases, not to the sugar or the phosphate backbone. The absorption spectrum of the HOP-DNA adduct indicates that the chromophore of HOP in the HOP-DNA covalent adduct seems to be similar to HOP itself since the features of the absorption spectra for both are virtually identical (Figure 3). This means that the chromophore is likely intact. Also, the strong fluorescence quenching for HOP upon HOP-DNA covalent adduct formation suggests that the pyrene ring of HOP is bonded to a nucleic acid base as seen in BPDE, whose fluorescence is also greatly quenched upon binding to guanine (64). Further study is needed to answer questions such as on which base and on what position of the base the covalent bond is formed. Another type of DNA damage induced by light and HOP is DNA single-strand cleavage. The processes leading to DNA cleavage appear to be through the excited singlet state of HOP and the generation of singlet oxygen and superoxide free radicals. Considering all the facts, we believe that HOP first reaches the excited singlet state by absorbing a photon. The singlet-state HOP is very energetic and can either react with DNA to form HOPDNA covalent adducts, with molecular oxygen (3O2), or be degraded. During these reactions, reactive intermediates such as singlet oxygen, superoxide, and HOP cation

592

Chem. Res. Toxicol., Vol. 13, No. 7, 2000

radicals may be generated. Both the ROS and HOP cation radicals should be capable of causing DNA cleavage and facilitate the formation of HOP-DNA adducts, as well as other forms of DNA damage, such as oxidation of guanine into 8-oxo- or 8-hydroxyguanine, depurination or depyrimidation, DNA-DNA strand cross-links, etc. Further work is needed to formulate a concise mechanism, to confirm other forms of DNA damage, and to find out cleavage sequence selectivity. In summary, the findings here present a possible light activation pathway for HOP. The covalent HOP-DNA adduct formation may especially present a new mechanism for the genotoxicity of HOP, or in general, for any PAHs that may form covalent adducts with DNA upon light activation. Our results with other PAHs reveal that they also can cause light-induced DNA cleavage, although their DNA cleavage potency is different from PAH to PAH (51). Resembling the reactions of benzo[a]pyrene and other PAHs which form DNA covalent adducts via the diol epoxide or one-electron oxidation intermediates after metabolic activation (3, 12, 13, 20), the light-induced reaction of HOP with DNA can produce DNA covalent adducts and cause other forms of DNA damage. These results indicate that in addition to enzymatic activation, light activation of PAHs may also be considered a potential toxicity source, especially since the DNA covalent adduct can be formed through light activation. Although PAH molecules present inside the human body may not be susceptible to light activation, those in the skin cells may be activated by light. It is known that absorption of PAHs through skin is a main pathway in addition to inhalation (2, 3, 6, 8, 9). Therefore, studies of light-induced toxicity of PAHs toward skin cells may be of interest.

Acknowledgment. This research was supported by the National Institutes of Health through generous grants (NIH-RCMI G12RR13459 and NIH-MBRS S06GM08047). We thank the Department of Energy for financial support under Contract DE-FG02-97ER62451 to the University of Georgia and Subcontract RR100-239/ 4891914 to Jackson State University. We thank Dr. J. B. Chaires from the University of Mississippi Medical Center for important suggestions concerning this project.

References (1) Baum, E. (1978) Occurrence and durveillance of polycyclic aromatic hydrocarbons. In Polycyclic Aromatic Hydrocarbons and Cancer (Gelboin, H., and Tso, T. O. T., Eds.) Vol. 1, pp 45-70, Academic Press, New York. (2) Connell, D. W., Hawker, D. W., Warne, M. J., and Vowles, P. P. (1997) Polycyclic aromatic hydrocarbons (PAHs). In Introduction into Environmental Chemistry (McCombs, K., and Starkweather, A. W., Eds.) pp 205-217, CRC Press, Boca Raton, FL. (3) Dipple, A. (1985) Polycyclic aromatic hydrocarbon carcinogenesis: an introduction. In Polycyclic Hydrocarbons and Carcinogenesis (Harvey, R. G., Ed.) ACS Symposium Series 283, pp 1-17, American Chemical Society, Washington, DC. (4) Conney, A. H. (1982) Induction of microsomal enzymes by foreign chemicals and carcinogenesis by polycyclic aromatic hydrocarbons: G. H. A. Clowes memorial lecture. Cancer Res. 42, 48754910. (5) Lesko, S. A. (1984) Chemical carcinogenesis: benzopyrene system. Methods Enzymol. 105, 539-550. (6) National Toxicology Program, U.S. Department of Health and Human Services (1998) Polycyclic aromatic hydrocarbons. In 8th Report on Carcinogens, pp 178-181, Integrated Laboratory Systems, Inc., Research Triangle Park, NC. (7) Warshawsky, D. (1999) Polycyclic aromatic hydrocarbons in carcinogenesis. Environ. Health Perspect. 107, 317-320.

Dong et al. (8) Talaska, G., Underwood, P., Maier, A., Lewtas, J., Rothman, N., and Jaeger, M. (1996) Polycyclic aromatic hydrocarbons (PAHs), nitro-PAHs and related environmental compounds: biological markers of exposure and effects. Environ. Health Perspect. 104, 701-908. (9) International Agency for Research on Cancer (1983) Polynuclear Aromatic Compounds. Part I: Chemical, Environmental and Experimental Data, Lyon, France. (10) U.S. Department of Health and Human Services, P. H. S., ATSDR (1995) Toxicological Profile for Polycyclic Aromatic Hydrocarbons (PAHs), Atlanta, GA. (11) 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. (12) RamaKrishna, N. V. S., Devanesan, P. D., Rogan, E. G., Cavalieri, E. L., Jeong, H., Jankowiak, R., and Small, G. J. (1992) Mechanism of metabolic activation of the potent carcinogen 7,12dimethylbenz[a]anthracene. Chem. Res. Toxicol. 5, 220-226. (13) Gelbroin, H. V. (1980) Benzo[a]pyrene metabolism, activation and carcinogenesis: role and regulation of mixed function oxidases and related enzymes. Physiol. Rev. 60, 1107-1166. (14) Geacintov, N. E., Cosman, M., Hingerty, B. E., Amin, S., Broyde, S., and Patel, D. J. (1997) NMR solution structure of stereoisomeric covalent polycyclic aromatic carcinogen-DNA adducts: principles, patterns, and diversity. Chem. Res. Toxicol. 10, 111146. (15) Harvey, R. G. (1991) Polycyclic Aromatic Hydrocarbons: Chemistry and Carcinogenicity, Cambridge University Press, Cambridge, U.K. (16) Penning, T. M., Burczynski, M. E., Hung, C. F., McCoull, K. D., Palackal, N. T., and Tsuruda, L. S. (1999) Dihydrodiol dehydrogenases and polycyclic aromatic hydrocarbon activation: generation of reactive and redox active o-quinones. Chem. Res. Toxicol. 12, 1-18. (17) Thakker, D. R., Yagi, H., Lu, A. Y. H., Levin, W., Conney, A. H., and Jerina, D. M. (1976) Metabolism of benzo[a]pyrene: conversion of trans-7,8-dihydroxy-7,8-dihydrobenzo[a]pyrene to highly mutagenic 7,8-diol-9,10-epoxides. Proc. Natl. Acad. Sci. U.S.A. 73, 3381-3385. (18) Devanesan, P. D., Higginbotham, S., Ariese, F., Jankoviak, R., Suh, M., Small, G. J., Cavalieri, E. L., and Rogan, E. G. (1996) Depurinating and stable benzo[a]pyrene-DNA adducts formed in isolated rat liver nuclei. Chem. Res. Toxicol. 9, 1113-1116. (19) Rogan, E. G., Devanesan, P. D., RamaKrishna, N. V. S., Higginbotham, S., Padmavathi, N. S., Chapman, K., Cavalieri, E. L., Jeong, H., Jankowiak, R., and Small, G. J. (1993) Identification and quantitation of benzo[a]pyrene-DNA adducts formed in mouse skin. Chem. Res. Toxicol. 6, 356-363. (20) Flowers, L., Ohinishi, S. T., and Penning, T. M. (1997) DNA strand scission by polycyclic aromatic hydrocarbon o-quinones: role of reactive oxygen species, Cu(II)/Cu(I) redox cycling, and o-semiquinone anion radicals. Biochemistry 36, 8640-8648. (21) Huang, X.-D., Krylov, S. N., Ren, L., McKonkey, B. J., Dixon, D. G., and Greenberg, B. M. (1997) Mechanistic quantitative structure-activity relationship model for the photoinduced toxicity of polycyclic aromatic hydrocarbons: II. An empirical model for the toxicity of 16 polycyclic aromatic hydrocarbons to the duckweek lemna gibba L. G-3. Environ. Toxicol. Chem. 16, 2296-2303. (22) Kagan, J., Tuveson, R. W., and Gong, H.-H. (1989) The lightdependent cytotoxicity of benzo[a]pyrene: effect on human erythrocytes, Escherichia coli cells, and Haemophilus influenzae transforming DNA. Mutat. Res. 216, 231-242. (23) Krylov, S. N., Huang, X.-D., Zeiler, L. F., Dixon, D. G., and Greenberg, B. M. (1997) Mechanistic quantitative structureactivity relationship model for the photoinduced toxicity of polycyclic aromatic hydrocarbons: I. Physical model based on chemical kinetics in a two-compartment system. Environ. Toxicol. Chem. 16, 2283-2295. (24) Mezey, P. G., Zimpel, Z., Warburton, P., Walker, P. D., Irvine, D. G., Huang, X.-D., Dixon, D. G., and Greenberg, B. M. (1998) Use of quantitative shape-activity relationship to model the photoinduced toxicity of polycyclic aromatic hydrocarbons: electron density shape features accurately predict toxicity. Environ. Toxicol. Chem 17, 1207-1215. (25) Pelletier, M. C., Burgess, R. M., Ho, K. T., Kuhn, A., McKinney, R. A., and Ryba, S. A. (1997) Phototoxicity of individual polycyclic aromatic hydrocarbons and petroleum to marine invertebrate larvae and juveniles. Environ. Toxicol. Chem. 16, 2190-2199. (26) Swartz, R. C., Ferraro, S. P., Lamberson, J. O., Cole, F. A., Ozretich, R. J., Boese, B. L., Schults, D. W., Behrenfeld, M., and Ankley, G. T. (1997) Photoactivation and toxicity of mixtures of

PAH Light-Induced DNA Damage and Phototoxicity

(27)

(28)

(29)

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

(39)

(40)

(41)

(42)

(43)

(44)

(45)

(46)

polycyclic aromatic hydrocarbon compounds in marine sediment. Environ. Toxicol. Chem. 16, 2151-2157. Santamaria, L., Giordano, G. G., Alfisi, M., and Cascione, F. (1966) Effects of light on 3,4-benzopyrene carcinogenesis. Nature 210, 824-825. Brooks, P., and Lawley, P. D. (1964) Evidence for the binding of polynuclear aromatic hydrocarbons to the nucleic acids of mouse skin: relation between carcinogenic power of hydrocarbons and their binding to deoxyribonucleic acid. Nature 202, 781-784. Blackburn, G. M., and Taussig, P. E. (1975) The photocarcinogenicity of anthracene: photochemical binding to deoxynucleic acids in tissue culture. Biochem. J. 149, 289-291. Blackburn, G. M., Fenwick, R. G., Lockwood, G., and Williams, G. M. (1977) Photoproducts from DNA pyrimidine bases and polycyclic aromatic hydrocarbons. Nucleic Acids Res. 4, 24872494. Striste, G. F., Martinez, E., Martinez, A. M., and Brake, R. J. (1980) Photo-induced reactions of benzo[a]pyrene with DNA in vitro. Cancer Res. 40, 245-252. Lesko, S. A., Ts’o, P. O. P., and Umans, R. S. (1969) Interaction of nucleic acids. V. Chemical linkage of 3,4-benzopyrene to deoxyribonucleic acids in aqueous solution. Biochemistry 8, 22912298. Ts’o, P. O. P., and Lu, P. (1964) Interaction of nucleic acids, II. Chemical linkage of the carcinogenic 3,4-benzopyrene to DNA induced by photoradiation. Proc. Natl. Acad. Sci. U.S.A. 51, 272280. Hoard, D. E., Ratliff, R. L., Bingham, J. M., and Striniste, G. F. (1981) Reaction induced in vitro between model DNA and benzo[a]pyrene by near-ultraviolet radiation. Chem.-Biol. Interact. 33, 179-194. Liu, Z., Lu, Y., Rosenstein, B., Lebwohl, M., and Wei, H. (1998) Benzo[a]pyrene enhances the formation of 8-hydroxy-2′-deoxyguanosine by ultraviolet A radiation in calf thymus DNA and human epidermoid carcinoma. Biochemistry 37, 10307-10312. Li, B., Mao, B., Liu, T.-M., Xu, J., Dourandin, A., Amin, S., and Geacintov, N. E. (1995) Laser pulse-induced photochemical strand cleavage of site-specifically and covalently modified (+)-antibenzo[a]pyrene diol epoxide-oligonucleotide adducts. Chem. Res. Toxicol. 8, 396-402. Camalier, R. F., Gantt, R., Price, F. M., Stephens, E. V., Baeck, A. E., Taylor, W. G., and Sanford, K. K. (1981) Effect of visible light on benzo[a]pyrene binding to DNA of cultured human skin epithelial cells. Cancer Res. 41, 1789-1793. White, G. L., Fu, P. P., and Heflich, R. H. (1985) Effect of nitrosubstitution on the light-mediated mutagenicity of polycyclic aromatic hydrocarbons in Samonella typhimurium TA 98. Mutat. Res. 144, 1-7. Jongeneelen, F. J. (1994) Biological monitoring of environmental exposure to polycyclic aromatic hydrocarbons: 1-hydroxypyrene in urine of people. Toxicol. Lett. 72, 205-211. Dor, F., Dab, W., Empereur-Bissonnet, P., and Zmirou, D. (1999) Validity of biomarkers in environmental health studies: the case of PAHs and benzenes. Crit. Rev. Toxicol. 29, 129-168. Merlo, F., Anderson, A., Weston, A., Pan, C.-F., Haugen, A., Valerio, F., Reggiardo, G., Fontana, V., Garte, S., Puntoni, R., and Abbondandolo, A. (1998) Urinary excretion of 1-hydroxypyrene as a marker for exposure to urban air levels of polycyclic aromatic hydrocarbons. Cancer Epidemiol., Biomarkers Prev. 7, 147-155. Godschalk, R. W. L., Ostertag, J. U., Moonen, E. J. C., Neumann, H. A. M., Kleinjans, J. C. S., and van Schooten, F. J. (1998) Aromatic DNA adducts in human white blood cells and skin after dermal application of coal tar. Cancer Epidemiol., Biomarkers Prev. 7, 767-773. Angerer, J., Mannschreck, C., and Gundel, J. (1997) Occupational exposure to polycyclic aromatic hydrocarbons in a graphiteelectrode producing plant: biological monitoring of 1-hydroxypyrene and monohydroxylated metabolites of phenanthrene. Int. Arch. Occup. Environ. Health 69, 323-331. Roggi, C., Minoia, C., Sciarra, G. F., Apostoli, P., Maccarini, L., Magnaghi, S., Cenni, A., Fonte, A., Nidasio, G. F., and Micoli, G. (1997) Urinary 1-hydroxypyrene as a marker of exposure to pyrene: an epidemiological survey on a general population group. Sci. Total Environ. 199, 247-254. Siwinska, E., Mielzynska, D., and Kwapulinski, J. (1998) Evaluation of intra and inter individual variation of urinary 1-hydroxypyrene, a biomarker of exposure to polycyclic aromatic hydrocarbons. Sci. Total Environ. 217, 175-183. Strickland, P., Kang, D., and Sithisarakul, P. (1996) Polycyclic aromatic hydrocarbon metabolite in urine as biomarkers of exposure and effect. Environ. Health Perspect. 104, 927-935.

Chem. Res. Toxicol., Vol. 13, No. 7, 2000 593 (47) Lambert, M., Kremer, S., and Anke, H. (1995) Antimicrobial, phytotoxic, nematicidal, cytotoxic, and mutagenic activities of 1-hydroxypyrene, the initial metabolite in pyrene metabolism by the basidiomycete Crinipellis stipitaria. Bull. Environ. Contam. Toxicol. 55, 251-257. (48) Hauser, B., Schrader, B. H., and Bahadir, M. (1997) Comparison of acute toxicity and genotoxicity concentrations of single compounds and waste elutriate using the Microtox/Mutatox test system. Ecotoxicol. Environ. Saf. 38, 227-231. (49) Jin, L., Tran, D. Q., Ide, C. F., McLachlan, J. A., and Arnold, S. F. (1997) Several synthetic chemicals inhibit progesterone receptor-mediated transactivation in yeast. Biochem. Biophys. Res. Commun. 233, 139-146. (50) Nakajima, D., Kojima, E., and Suzuki, S. (1996) Presence of 1-hydroxypyrene conjugates in woody plant leaves and seasonal changes in their concentrations. J. Environ. Sci. Technol. 30, 1675. (51) Dong, S., Hwang, H.-M., Harrison, C., Holloway, L., Shi, X., and Yu, H. (2000) UVA light-induced DNA cleavage by selected polycyclic aromatic hydrocarbons. Bull. Environ. Contam. Toxicol. 64 (4), 467-474. (52) Ciulla, T. A., Van Camp, J. R., Rosenfeld, E., and Kochevar, I. E. (1989) Photosensitization of single strand breaks in pBR322 DNA by rose bengal. Photochem. Photobiol. 49, 293-298. (53) McGhee, J. D., and von Hippel, P. H. (1974) Theoretical aspects of DNA-protein interactions: cooperative and non-cooperative binding of large ligands to a one-dimensional homogeneous lattice. J. Mol. Biol. 86, 469-489. (54) Chaires, J. B., Priebe, W., Graves, D. E., and Burke, T. G. (1993) Dissection of the free energy of anthracycline antibiotic binding to DNA: electrostatic contributions. J. Am. Chem. Soc. 115, 5360-5364. (55) Wadkins, R. M., and Graves, D. E. (1989) Thermodynamics of the interaction of m-AMSA and o-AMSA with nucleic acids: influence of ionic strength and DNA base composition. Nucleic Acids Res. 17, 9933-9946. (56) Price, H. L., Fetzer, S. M., and LeBreton, P. R. (1990) Evidence for nonintercalative complexes formed from the reversible binding of benzo[a]pyrene metabolites to closed-circular, single-stranded M13mp19 DNA. Biomed. Biophys. Res. Commun. 168, 10951102. (57) Meehan, T., Gamper, H., and Becker, J. F. (1982) Characterization of reversible, physical binding of benzo[a]pyrene derivatives to DNA. J. Biol. Chem. 257, 10479-10485. (58) Geacintov, N. E., Zinger, D., Ibanez, V., Santella, R., Grunberger, D., and Harvey, R. G. (1987) Properties of covalent benzo[a]pyrene diol epoxide-DNA adducts investigated by fluorescence technique. Carcinogenesis 8, 925-935. (59) Nolan, S. J., Vyas, R. R., Hingerty, B. E., Ellis, S., Broyde, S., Shapiro, R., and Basu, A. K. (1996) Solution properties and computational analysis of an oligodeoxynucleotide containing N-(deoxyguanosin-8-yl)-1-aminopyrene. Carcinogenesis 17, 133144. (60) Leng, F., Savkur, R., Kokt, I., Przewloka, T., Priebe, W., and Chaires, J. B. (1996) Base specific and regioselective chemical cross-linking of Daunorubicin to DNA. J. Am. Chem. Soc. 118, 4731-4738. (61) Leng, F., Graves, D., and Chaires, J. B. (1998) Chemical crosslinking of ethidium to DNA by glyoxal. Biochim. Biophys. Acta 1442, 71-81. (62) Foote, C. S. (1979) Detection of singlet oxygen in complex systems: a critique. In Biochemical and Clinical Aspects of Oxygen (Caughey, W. S., Ed.) pp 601-626, Academic Press, New York. (63) Williams, R. M., Glinka, T., Flanigan, M. E., Gallegos, R., Coffman, H., and Pei, D. (1992) Cannizaro-based O2-dependent cleavage of DNA by quinocarcin. J. Am. Chem. Soc. 114, 733740. (64) Shafirovich, V. Y., Levin, P. P., Kuzmin, V. A., Thorgeirsson, T. E., Kliger, D. S., and Geacintov, N. E. (1994) Photoinduced electron transfer and enhanced triplet yields in benzo[a]pyrene derivative-nucleic acid complexes and covalent adducts. J. Am. Chem. Soc. 116, 63-72. (65) Miller, J. C., Meek, J. S., and Strickler, S. J. (1977) Heavy atom effects on the triplet lifetimes of naphthalene and phenanthrene. J. Am. Chem. Soc. 99, 6175-6179.

TX990199X