Enhancement of 2'-Deoxyguanosine Hydroxylation and DNA Damage

Sep 13, 2000 - Enhancement of 2'-Deoxyguanosine Hydroxylation and DNA Damage by Coal and Oil Fly Ash in Relation to Particulate Metal Content and Avai...
58 downloads 12 Views 89KB Size
Download PDF
Chem. Res. Toxicol. 2000, 13, 1011-1019

1011

Enhancement of 2′-Deoxyguanosine Hydroxylation and DNA Damage by Coal and Oil Fly Ash in Relation to Particulate Metal Content and Availability Agasanur K. Prahalad,† Jeff Inmon,‡ Andrew J. Ghio,‡ and Jane E. Gallagher*,‡ Curriculum in Toxicology, University of North Carolina, Chapel Hill, North Carolina 27599, and Human Studies Division, National Health and Environmental Effects Research Laboratory, U.S. Environmental Protection Agency, Research Triangle Park, North Carolina 27711 Received May 17, 2000

Epidemiologic studies have shown causal relationships between air pollution particles and adverse health effects in susceptible subpopulations. Fly ash particles (containing water-soluble and insoluble metals) are a component of ambient air particulate pollution and may contribute to particulate-induced health effects. Some of the pathological effects after inhalation of the particles may be due to reactive oxygen species (ROS) produced by metal-catalyzed reactions. In this investigation, we analyzed emission source particulates oil fly ash (OFA) and coal fly ash (CFA) for metal content and solubility in relation to their ability to induce 2′-deoxyguanosine (dG) hydroxylation and DNA damage as measured by 8-oxo-dG formation by HPLC/UVelectrochemical detection (ECD). Water-soluble vanadium and nickel were present at the highest concentrations, and iron was present in trace amounts in OFA (5.1% V, 1.0% Ni, and 0.4% Fe by weight). In contrast, CFA comprised mostly of water-insoluble aluminosilicates and iron (9.2% Al, 12.2% Si, and 2.8% Fe by weight). As a first approach to gain insight into the mode of action of these particulates, we examined metal species-catalyzed kinetics of dG hydroxylation. Metal species at a concentration of 0.1 mM in the incubation mixture containing 0.1 mM dG under ambient air at room temperature catalyzed maximum 8-oxo-dG formation at 15 min with yields ranging from 0.05 to 0.17%, decreasing in the following order: vanadium(IV) > iron(II) > vanadium(V) > iron(III) g nickel(II). Insoluble Fe(III) oxide (Fe2O3) under similar conditions had no effect. Consistent with these results, OFA rich in vanadium and nickel concentrations showed a dose-dependent increase in the level of dG hydroxylation to 8-oxo-dG formation at particulate concentrations of 0.1-1 mg/mL (p < 0.05). In contrast, CFA with high concentrations of aluminosilicates and iron did not result in a significant increase in the level of 8-oxo-dG over that of the control, i.e., dG (p > 0.05). DMSO, a •OH scavenger, inhibited OFA-induced 8-oxo-dG formation, and metal ion chelators, deferoxamine (DFX), DTPA, and ferrozine blocked OFA-induced 8-oxo-dG formation. OFA and CFA induced 8-oxodG formation in a pattern similar to that observed for dG hydroxylation when calf thymus DNA was used as a substrate. Treatment of OFA particles with DFX before reacting with DNA or addition of a catalase in the incubation mixture significantly suppressed 8-oxo-dG formation (p < 0.05). These results suggest that metal availability, but not the concentration of metals present in CFA and OFA, is critical in mediating molecular oxygen-dependent dG hydroxylation and DNA base damage.

Introduction Epidemiologic studies have demonstrated significant associations between exposures to ambient air pollution particles and adverse health effects in individuals with compromised respiratory systems (1-4). While biochemical mechanisms for particulate-induced health effects remain unclear (5), it has been hypothesized that such adverse health effects from particulate exposure can result from metal-mediated generation of reactive oxygen * To whom correspondence should be addressed: Human Studies Division, MD-58C, National Health and Environmental Effects Research Laboratory, U.S. EPA, Research Triangle Park, NC 27711. Phone: (919) 966-0638. Fax: (919) 966-0655. E-mail: gallagher.jane@ epamail.epa.gov. † University of North Carolina. ‡ U.S. Environmental Protection Agency.

species (ROS)1 (6, 7). Particulate air pollution contains transition metals in different proportions, form, and oxidation state, which can catalyze the reduction of molecular oxygen, generating ROS which exert a variety of biochemical effects (8, 9). Oil and coal fly ash are emission source particulate air pollutants. In rats, intratracheal instillation of these particulates induces an acute pulmonary injury charac1 Abbreviations: 8-oxo-dG, 7,8-dihydro-8-oxo-2′-deoxyguanosine; dG, 2′-deoxyguanosine; dA, 2′-deoxyadenosine; dC, 2′-deoxycytidine; T, thymidine; DFX, deferoxamine mesylate; DTPA, diethylenetriaminepentaacetic acid; ferrozine, 3-(2-pyridyl)-5,6-bis(4-phenylsulfonic acid)-1,2,4-triazine; DMSO, dimethyl sulfoxide; ROS, reactive oxygen species; CFA, coal combustion fly ash; OFA, oil combustion fly ash; BHT, butylated hydroxytoluene; SOD, superoxide dismutase; CAT, catalase; XRF, energy-dispersive X-ray fluorescence; HPLC-EC, highperformance liquid chromatography-electrochemical detection.

10.1021/tx000110j CCC: $19.00 © 2000 American Chemical Society Published on Web 09/13/2000

1012

Chem. Res. Toxicol., Vol. 13, No. 10, 2000

terized by neutrophilic alveolitis, airway hyperreactivity, and an increased susceptibility to microbial infections (10). The pulmonary injury induced by oil fly ash exposure is correlated with its high content of transition metals such as V and Ni, while coal fly ash-associated injury is related to its Fe content (10). Vanadium, nickel, and their derivatives have been shown to exert potent toxic effects in a wide variety of biological systems (9). Epidemiologic studies have shown a correlation between V exposure and the incidence of lung cancer in humans (11-13). Vanadium predominantly exists in either V(IV) or vanadyl and V(V) or vanadate ions under physiological conditions (14). Vanadate has been shown to induce forward mutations and DNA protein cross-links (14). Nickel and its derivative in experimental animals and humans manifest as a lung and sinonasal carcinogen (15, 16). While iron is essential for life, excessive exposure to this metal becomes a risk factor in many human diseases, including cancer (17, 18). An increasing interest has developed with regard to the role of metals in nucleic acid damage (19). However, most of the emphasis has been placed on Fenton-driven Fe and Cu reactions (19). While the biochemical mechanism of V and Ni carcinogenicity and toxicity is still not fully understood, studies have indicated that V- and Ni-mediated generation of ROS may play an important role (9). We have previously demonstrated the cellular production of oxygen-based radicals upon exposure to a wide range of ambient air particles as measured by luminol-induced chemiluminescence in human blood leukocytes (20). Although the assay is simple and sensitive for cellular production of oxygen metabolites, it is not specific and will not discriminate between the individual oxygen species (21), and accordingly, the nature of any ROS being formed cannot be ascertained. However, it has been demonstrated that hydroxylation at the C-8 position of 2′-deoxyguanosine (dG) residues in DNA by either hydroxyl radical (•OH) or a species with a similar reactivity results in the formation of 8-oxo-2′-deoxyguanosine (8-oxo-dG), providing a strong indication of •OH production (22). The purpose of this work was to determine the ability of oil fly ash (OFA) and coal fly ash (CFA) to hydroxylate the C-8 position of the guanine residue within the corresponding 2′-deoxyribonucleoside and calf thymus DNA. The hydroxylation product, 8-oxo-dG, was assessed by using a selective and sensitive HPLC electrochemical technique. In addition, it was of interest to determine how the solubility and metal content of the particles and protective effects of different metal ion chelators, •OH scavenger, and antioxidant enzymes affect 8-oxo-dG formation. Although 8-oxo-dG formation is one measure of oxidative damage, other transition metal ion-mediated DNA oxidation produces a wide spectrum of damage, including base lesions, oligonucleotide strand breaks, abasic sites, and DNA-protein cross-links (22, 23). In this investigation, we examined 8-oxo-dG formation because this DNA lesion is a widely accepted marker of oxidative damage.

Materials and Methods Caution: Nickel and vanadium compounds are toxic, mutagenic, and suspected human carcinogens and should be handled with extreme caution. Biochemicals and Reagents. All chemicals and reagents were analytical or enzyme grade unless otherwise mentioned.

Prahalad et al. 2′-Deoxyguanosine (dG), 2′-deoxyadenosine (dA), 2′-deoxycytidine (dC), thymidine (T), calf thymus DNA, butylated hydroxytoluene (BHT), ascorbic acid, sodium acetate, tris(hydroxymethyl)aminomethane (Tris), mono- and dibasic sodium phosphate, EDTA, proteinase K, RNase A, RNase T1, catalase, superoxide dismutase (SOD), ethyl alcohol, sodium chloride, deferoxamine mesylate (DFX), diethylenetriaminepentaacetic acid (DTPA), 3-(2-pyridyl)-5,6-bis(4-phenylsulfonic acid)-1,2,4-triazine (ferrozine), dimethyl sulfoxide (DMSO), ferrous sulfate (FeSO4‚ 7H2O), ferric sulfate [Fe2(SO4)3], and ammonium vanadate (NH4VO3) as a source of vanadium(V) were purchased from Sigma Chemical Co. (St. Louis, MO). Sodium citrate dihydrate was bought from Mallinckrodt (Paris, KY). Iron(III) oxide (Fe2O3), vanadyl sulfate (VOSO4‚4H2O) as a source of vanadium (IV), and nickel(II) sulfate (NiSO4‚6H2O) were obtained from Johnson Matthey Co. (Ward Hill, MA). Nuclease P1 and alkaline phosphatase were from Boehringer Mannheim (Indianapolis, IN). HPLC grade methanol was purchased from Fisher Scientific (Fairlawn, NJ). All glassware was rinsed with a mixture of sulfuric and nitric acids (1:1) to eliminate trace metals, followed by a wash with glass-distilled water, dried, and autoclaved prior to use. Particle Source. OFA is an oil combustion fly ash collected from an electrostatic precipitator (collection temperature of 340 °C) at an eastern U.S. power plant (Niagara, NY). The particulate has a mass median aerodynamic diameter of 3.18 ( 0.44 µm. Coal combustion fly ash (CFA) is a conventional coal combustion fly ash obtained from a western U.S. power plant burning western coal. This dust has a mass median aerodynamic diameter of 4.25 ( 0.35 µm. The physicochemical characteristics and elemental composition of the particulates utilized in this studies have been described in detail previously (10, 20). Synthesis of the 8-Oxo-dG Standard. Standard 8-oxodG was sythesized by Udenfriend’s hydroxylating system as described by Kasai and Nishimura (24) and purified by HPLC on a reverse phase column as described below. Briefly, hydroxylation of dG at the C-8 position was carried out by sequential addition of 25 µL of 0.1 M dG, 14 µL of 1 M ascorbic acid, 6.5 µL of 1 M EDTA, and 13 µL of 0.13 M FeSO4 (all freshly prepared) to 0.942 µL of 0.1 M sodium phosphate buffer (pH 6.8) and incubation for up to 15 min at 37 °C in the dark with vigorous shaking. Following the incubation, aliquots of the reaction mixture were analyzed by reverse phase HPLC using a 0.46 cm × 25 cm Beckman Ultrasphere ODS column protected by a 0.46 cm × 4.5 cm Ultrasphere ODS guard column (Beckman Instruments, Fullerton, CA). The column was eluted isocratically at a flow rate of 1 mL/min with a 1:9 (v/v) methanol/ water mixture. The eluates were monitored at 254 nm. Fractions containing 8-oxo-dG, which eluted around 28 min, were manually collected, lyophilized, and used as a standard. The yield of 8-oxo-dG formation was estimated to be approximately 10%, as judged by the HPLC/UV peak. The identity of 8-oxo-dG was further confirmed by UV-visible spectroscopy, showing λmax ) 247 and 293 nm in distilled H2O in agreement with the reported values (25), and also coelution with authentic standard 8-oxo-dG obtained from ESA, Inc. (Chelmsford, MA). Metal-Catalyzed Hydroxylation of dG to 8-Oxo-dG. The time course of conversion of dG to 8-oxo-dG formation by metal ions was studied by incubating 0.1 mM sulfate or ammonium salts of metal species [V(IV), V(V), Fe(II), Fe(III), or Ni(II)], or Fe(III) oxide, with 0.1 mM dG in 1 mL of 100 mM phosphate buffer (pH 7.4). The incubation mixture was kept open in the dark at room temperature under ambient air to allow air exchange and analyzed immediately and thereafter at 2, 5, 10, 15, or 20 min intervals for 8-oxo-dG formation by HPLC-coupled UV-EC detection as described below. The experiments were also performed in the presence of different metal ion chelators such as DFX, DTPA, or ferrozine at concentrations of 0.2 mM in the reaction mixture. Particulate-Mediated Hydroxylation of dG to 8-Oxo-dG. dG was incubated at ambient temperature for 15 min in the dark with different concentrations of OFA or CFA alone or in

Particulate-Induced Oxidative DNA Base Damage combination with different metal ion chelators in phosphatebuffered suspension in 2 mL capacity mircofuge tubes. The tubes were vortexed for 30 s and kept open to allow air exchange at room temperature. The concentrations of reactants in the incubation mixture that were studied were as follows: 0.1 mM dG, 0.1-1 mg/mL particulate (OFA or CFA), 0.2 mM metal ion chelator DFX, ferrozine, or DTPA, and 2 M (5%) •OH scavenger DMSO in a total volume of 1 mL of 100 mM phosphate buffer (pH 7.4). Following incubation, the reaction mixture was filtered through a SPIN-X 0.45 mm nylon centrifuge tube filter in a 2 mL tube (Corning Costar Corp.) and the resulting filtrate was analyzed for dG and 8-oxo-dG by HPLC coupled with UV-EC detection as described below. Typically, injection volumes of the filtrate did not exceed 20 µL. Particulate-Induced Oxidation of Isolated DNA. Calf thymus DNA was freed from RNA and proteins by digestion with RNases and proteinase K, respectively, followed by solvent extractions and ethanol precipitation by a modified procedure of Marmur (26), without the use of phenol during DNA extraction. Furthermore, 30 mM BHT (27) was added as an antioxidant prior to the solvent extraction. DNA was dissolved in distilled water, quickly frozen in liquid nitrogen, and stored at -70 °C until it was required. Particulates (0.1-1 mg/mL) were incubated with DNA (0.5 mg/mL) for 15 min in the presence or absence of metal ion chelator DFX (0.2 mM) and antioxidant enzyme catalase (2000 units/mL) or SOD (30 units/mL) in a total volume of 500 mL of 100 mM phosphate buffer (pH 7.4) in 2 mL capacity microfuge tubes. The tubes were vortexed for 30 s and kept open to allow air exchange at room temperature. Following incubation, the particulates were removed by centrifugation at 13000g for 3 min at 4 °C. Subsequently, 50 µL of 3 M sodium acetate and 2 volumes of cold ethanol were added to the supernatant, and the mixture was cooled at -70 °C for 15 min prior to centrifugation for 5 min. The resulting oxidized DNA precipitate was then redissolved in 500 µL of distilled water for subsequent enzymatic hydrolysis. Enzymatic Hydrolysis of DNA. Approximately 50 µg of DNA was removed and brought to a volume of 100 µL in 0.01 M Tris-HCl (pH 7.0). Five microliters of 0.5 M sodium acetate (pH 5.1) and 2.5 units of nuclease P1 (600 units/mL) were added, and the mixture was incubated at 37 °C for 30 min in the dark. After addition of 40 µL of 0.4 M Tris-HCl (pH 7.5) and 2 units of alkaline phosphatase, the mixture was further incubated for 1 h at 37 °C. The resulting hydrolysates were centrifuged at 13000g for 10 min at 4 °C, and the supernatants were stored on ice for no more than 6-8 h prior to analysis by HPLC/UVelectrochemical detection (ECD). Determination of dG and 8-Oxo-dG by HPLC/UV-ECD. The HPLC analytical system consisted of a model 500 solvent delivery dual piston pump (ESA, Inc., Chelmsford, MA) equipped with a Rheodyne loop injector, model 9125 (Cotati, CA), an UV/ Vis model 520 detector, and a model 5200 Coulochem II multielectrode ECD system (both from ESA Inc.). An ESA system RS232 interface module was connected directly to both the computer and to the HPLC pumps that were in turn interfaced to the UV-Vis and a Coulochem multielectrode EC detector configured in series. Three modular potentiostats were connected to a model 5020 guard cell positioned between the pump and the injector, a model 5021 conditioning cell, and a model 5010 dual electrode analytical cell. The guard cell includes a single large porous graphite working electrode with counter and reference electrodes. The potential of the guard cell was set at 450 mV to remove inherent electroactive impurities from the mobile phase, thereby lowering background currents. The potential of the conditioning cell and electrode one of the dual electrode analytical cell was set at 100 and 150 mV, respectively (used as screens), and located after the column and UV-Vis detector. Screen oxidation occurs at these electrodes at the set potentials such that some sample interferences, if any, are oxidized and therefore do not interfere with the analyte of our interest, 8-oxo-dG, which is subsequently analyzed at the second electrode of the sequential dual-electrode analytical cell set at

Chem. Res. Toxicol., Vol. 13, No. 10, 2000 1013 350 mV as optimized by maximum current response exhibited by 8-oxo-dG in the current-voltage curve. The analytical 0.46 cm × 25 cm Ultrasphere ODS column protected by 0.46 cm × 4.5 cm ODS guard column (Beckman Instruments, Fullerton, CA) was used for dG and 8-oxo-dG determinations in samples. The eluant was 4-5% aqueous methanol containing 12.5 mM sodium citrate and 25 mM sodium acetate (pH 5.1) at a flow rate of 1 mL/min. The HPLC mobile phase was filtered prior to use through a 0.45 µm HV Millipore filter (Milford, MA) for degassing purposes. Statistical Analysis. The software package Prism, version 2.01, from Graph Pad (San Diego, CA) was used for the statistical analysis. The data are expressed as means ( SD (n ) 3). A two-tailed paired Student’s t test at the 95% confidence interval was used for pairwise comparisons. Statistically significant differences are reported when p < 0.05.

Results Particulate Chemistry. Concentrations and the solubility and elemental composition of the OFA and CFA as determined by energy-dispersive X-ray fluorescence (XRF) analyses are described in detail elsewhere (20). Briefly, the percent of sample mass accounted for by XRF-detected elements in CFA and OFA constitutes approximately 28 and 13%, respectively. The majority of the sample mass unaccounted for by XRF consists of C, N, O, water, or organic constituents. The water-soluble elemental masses in CFA and OFA were ∼2 and 82%, respectively. CFA was found to be comprised mostly of water-insoluble aluminosilicates and Fe (12.2% Si, 9.2% Al, and 2.8% Fe by weight). In contrast, OFA contains high concentrations of water-soluble V and Ni, but Fe was present in trace amounts (5.1% V, 1.0% Ni, and 0.4% Fe by weight). The soluble metals in the particulates are likely to exist as sulfates and/or ammonium salts. The metals retained in the insoluble component are probably mineral oxides (7, 10). Metal Species-Mediated Hydroxylation of dG at the C-8 Position. The CFA and OFA used in this study contain transition metals in different proportions and form. In what form (soluble vs insoluble) these metals exist in particulates determines the biological activity of the inhaled particulates. We compared different metal species-mediated and particulate-induced C-8 hydroxylation of dG in relation to particulate metal content and solubility. Initially, a time course study was carried out to determine the optimum period of exposure of dG to the hydroxylating action of several metal ion species at concentrations corresponding to approximately 5 µg of metal/mL of reaction mixtures. This concentration was chosen on the basis of the concentrations of major metals Ni, V, and Fe associated with the OFA or CFA ranging between 1 and 5 µg per 100 µg of particulate per milliliter of incubation mixture. All metal species induced hydroxylation of dG to 8-oxo-dG, and the level was maximum around 15 min as shown in Figure 1. A similar time dependence was noted for V(IV) or vanadyl ion-mediated conversion of dG to 8-oxo-dG (28). Figure 2 illustrates representative HPLC/UV-EC profiles obtained from the reaction of freshly prepared 0.1 mM dG and 0.1 mM metal species in 100 mM phosphate buffer. The profiles compare UV (lower trace) and EC detection (upper trace). Under the conditions described here, 8-oxodG elutes around 20-23 min as a well-resolved peak. The elution time was confirmed by the spiking of the reaction mixture with purified standard 8-oxo-dG. Variations in

1014

Chem. Res. Toxicol., Vol. 13, No. 10, 2000

Figure 1. Time courses of conversion of dG to 8-oxo-dG. Metal ions (0.1 mM) were incubated in 100 mM phosphate buffer (pH 7.4) with 0.1 mM dG. The incubation mixture was kept open in the dark at room temperature under ambient air to allow air exchange and analyzed for 8-oxo-dG formation by HPLC coupled with UV-EC detection as described in Materials and Methods. The data points are the mean of duplicate analysis.

Prahalad et al.

Figure 3. Relative percentage yields of 8-oxo-dG formation from dG hydroxylation by metal ions. Metal ions (0.1 mM) in aqueous buffered solutions (pH 7.4) were incubated with dG (0.1 mM) for 15 min at room temperature under ambient air. An aliquot of the reaction mixture was analyzed for dG and 8-oxodG formation by HPLC/UV-ECD as described in Materials and Methods. The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. Table 1. Inhibition of Metal Ion-Mediated Hydroxylation of dG to 8-Oxo-dGa by Metal Ion Chelators

Figure 2. Representative HPLC/UV-EC profiles obtained from the reaction of freshly prepared metal ions (0.1 mM) and dG (0.1 mM) in 100 mM phosphate buffer (pH 7.4): lower profiles, UV (A253); upper profiles, EC detector response. For EC clarity, only the eluant after 10 min has been shown in the upper trace. The sample was injected after incubation for 15 min at room temperature under ambient air. See Materials and Methods for HPLC conditions. Under the conditions described herein, dG elutes between 12 and 15 min and 8-oxo-dG around 20-23 min.

the retention time occurred for dG and 8-oxo-dG due to fluctuations in the ambient laboratory temperature. 2′-Deoxyguanosine alone in a phosphate buffer produced 1.9 8-oxo-dG molecules/105 dG molecules. Storage of dG solution frozen and left in the dark over time did not increase the amount of 8-oxo-dG. Incubation of dG with metal ions significantly increased the level of 8-oxo-dG formation with yields ranging from 0.05 to 0.17% and decreased in the following order: V(IV) > Fe(II) > V(V) > Fe(III) g Ni(II). Incubation under argon prevented the hydroxylation, indicating an important role of molecular oxygen in 8-oxo-dG formation. Similar results were noted for vanadyl and Ni(II)-mediated hydroxylation of dG (28, 29). Insoluble Fe2O3 had no effect (Figure 3). Effects of Metal Ion Chelators on Metal SpeciesMediated dG Hydroxylation. Table 1 shows the effects of metal ion chelators on hydroxylation of dG to 8-oxodG by different metal species. These chelators are often used to determine whether metal ions are involved in the production of radicals. Preincubation of metals in

incubation mixtureb

8-oxo-dG/105 dGc

dG alone dG and V(IV) dG, V(IV), and DFX dG, V(IV), and DTPA dG, V(IV), and ferrozine dG and V(V) dG, V(V), and DFX dG, V(V), and DTPA dG, V(V), and ferrozine dG and Fe(II) dG, Fe(II), and DFX dG, Fe(II), and DTPA dG, Fe(II), and ferrozine dG and Fe(III) dG, Fe(III), and DTPA dG and Ni(II) dG, Ni(II), and DFX dG, Ni(II), and DTPA dG, Ni(II), and ferrozine

1.9 ( 0.15 169.1 ( 21.94 42.0 ( 5.60 (75.2)d 18.3 ( 2.80 (89.2) 53.5 ( 14.45 (68.4) 71.3 ( 7.35 24.3 ( 2.20 (65.9) 11.0 ( 0.10 (84.6) 18.1 ( 1.80 (74.6) 94.1 ( 6.05 9.7 ( 0.25 (89.7) 23.3 ( 0.90 (75.2) 21.4 ( 1.35 (77.3) 47.0 ( 1.00 3.8 ( 0.38 (91.9) 52.5 ( 4.55 17.0 ( 0.65 (67.6) 8.1 ( 0.45 (84.6) 22.3 ( 0.55 (57.5)

a Determined by HPLC-EC analysis. b Concentrations of reactants in the incubation mixture were as follows: 0.1 mM dG, 0.1 mM metal ions, and 0.2 mM metal ion chelators. The reaction mixtures were incubated in 100 mM phosphate buffer (pH 7.4) at room temperature for 15 min under ambient air. c The data are presented as the means ( the SD (n ) 3). d Values in parentheses indicate the percent inhibition of 8-oxo-dG formation by metal ion chelators.

0.2 mM chelator for 2 h before addition of dG inhibited 8-oxo-dG formation (Table 1). These findings show that the active coordination sites of the metal ions may be blocked by the chelators which render them redox inactive, confirming the importance of available or the catalytically active form of metals in the production of ROS and their subsequent effect on 8-oxo-dG formation. Particulate-Induced Hydroxylation of dG at the C-8 Position. The results of the comparative study on the C-8 hydroxylation of the purine moiety of dG by CFA and OFA are shown in Table 2. Incubation of dG (0.1 mM) with OFA (1 mg/mL) rich in water-soluble V and Ni resulted in a significant increase in the extent of 8-oxodG formation over the control dG (p < 0.05). On the other hand, CFA (1 mg/mL) containing a high concentration of insoluble Fe exhibited only a weak or no hydroxylating

Particulate-Induced Oxidative DNA Base Damage Table 2. Particulate (CFA and OFA)-Induced Hydroxylation of dG to 8-Oxo-dGa

Chem. Res. Toxicol., Vol. 13, No. 10, 2000 1015 Table 3. Effect of Metal Ion Chelators and DMSO on Particulate-Induced Hydroxylation of dG to 8-Oxo-dG

particulate

8-oxo-dG/105 dGb

incubation mixturea

8-oxo-dG/105 dGb

control (dG) CFA and dG OFA and dG

1.95 ( 0.15 2.03 ( 0.39 32.60 ( 2.40c

OFA and dG OFA, dG, and ferrozine OFA, dG, and DTPA OFA, dG, and DFX OFA, dG, and DMSO CFA and dG CFA, dG, and ferrozine CFA, dG, and DTPA CFA, dG, and DFX CFA, dG, and DMSO

32.60 ( 2.40 27.50 ( 1.50 19.00 ( 1.00c 10.89 ( 0.27c 3.20 ( 0.30c 2.03 ( 0.39 1.96 ( 0.31 2.16 ( 0.16 1.94 ( 0.07 2.15 ( 0.15

a Particulates (1 mg/mL) were incubated in 100 mM phosphate buffer (pH 7.4) with 0.1 mM dG for 15 min at room temperature under ambient air. dG and 8-oxo-dG were determined by HPLC analysis with UV-EC detections as described in Materials and Methods. b The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. c Significant increase relative to control dG (p < 0.05).

a Particulates (1 mg/mL) were preincubated with or without metal ion chelators (0.2 mM) or DMSO, 2 M (5%) before addition to dG (0.1 mM) in phosphate buffer (pH 7.4). Incubation was carried out for 15 min at room temperature under ambient air. Following incubation, dG and 8-oxo-dG levels were measured by HPLC/UV-EC analysis as described in Materials and Methods. b The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. c Significant inhibition of 8-oxo-dG (p < 0.05).

Figure 4. Effect of OFA concentrations on the efficiency of maximal dG hydroxylation. OFA particulates at the indicated concentrations were incubated with 0.1 mM dG in phosphate buffer (pH 7.4) for 15 min at room temperature under ambient air. Following incubation, dG and 8-oxo-dG levels were measured by HPLC/UV-EC analysis as described in Materials and Methods. The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. The absence of an error bar indicates that the SD is contained within the symbol.

capability as measured by the level of 8-oxo-dG formation(p > 0.05). Figure 4 illustrates the dose-dependent increase in the extent of hydroxylation of dG to 8-oxodG by OFA at incubation mixture concentrations ranging from 0.1 to 1 mg/mL. It should be mentioned that the low efficiency with which CFA hydroxylated dG cannot be explained in terms of complexation of nucleoside by this material. This was inferred from the observation that the addition of insoluble hematite, i.e., Fe2O3 or CFA, in suspension to the aqueous solution containing trace amounts of 8-oxo-dG did not show any significant reduction in the quantitative response on the HPLC-EC detection. Effects of Metal Ion Chelators and DMSO on Particulate-Induced dG Hydroxylation. To confirm whether available or insoluble metal components associated with the emission source particulates are responsible for hydroxylation of dG to 8-oxo-dG, the metal ion chelators were used. As shown in Table 3, incubation of OFA at a concentration of 1 mg/mL in 0.2 mM metal ion chelator such as DFX, DTPA, or ferrozine for 2 h before the addition of 0.1 mM dG in 100 mM phosphate buffer (pH 7.4) resulted in inhibition of 8-oxo-dG formation, with deferoxamine showing the highest level of inhibition followed by DTPA and ferrozine. Furthermore, DMSO, a well-established •OH scavenger at a concentration of 2 M (5%) in the incubation mixture, dramatically suppressed 8-oxo-dG formation. In contrast, CFA-induced 8-oxo-dG formation was largely unaffected by the treat-

Figure 5. Representative HPLC/UV-EC profiles of calf thymus DNA hydrolysates (A) or DNA treated with OFA (1 mg/mL). The profiles are a comparison of the UV (lower trace) and EC detection (upper trace). For EC clarity, only the eluant after 10 min has been shown in the upper trace. The 8-oxo-dG elutes around 25-26 min in each case. See Materials and Methods for HPLC conditions.

ment of the particles either with metal ion chelators or by DMSO treatment (Table 3). Particulate-Induced 8-Oxo-dG Formation within Calf Thymus DNA. The induction of 8-oxo-dG was assessed in calf thymus DNA in buffered solution containing suspensions of emission source particulates treated with or without metal ion chelator deferoxamine or the antioxidant enzymes catalase and superoxide dismutase (SOD). Figure 5 illustrates representative HPLC/UVEC profiles of calf thymus DNA hydrolysates (panel A) or DNA treated with OFA (panel B). The 8-oxo-dG elutes immediately after T and before dA at approximately 2526 min. The elution time was confirmed by the spiking of DNA hydrolysates with purified standard 8-oxo-dG in

1016

Chem. Res. Toxicol., Vol. 13, No. 10, 2000

Table 4. Particulate-Induced 8-Oxo-dG Formation in Calf Thymus DNAa particulate

8-oxo-dG/105 dGb

control (DNA) CFA and DNA OFA and DNA

7.20 ( 0.26 9.75 ( 1.67 26.49 ( 0.31c

a Particulates (1 mg/mL) were incubated in 100 mM phosphate buffer (pH 7.4) with calf thymus DNA for 15 min at room temperature under ambient air in a total volume of 0.5 mL, followed by enzymatic hydrolysis of DNA to nucleosides. The DNA hydrolysates were analyzed for dG and 8-oxo-dG levels by HPLC analysis with UV-EC detections as described in Materials and Methods. b The results are expressed as means ( the SD (n ) 3). c Significant induction of 8-oxo-dG relative to control DNA (p < 0.05).

Prahalad et al. Table 5. Effect of Catalase, SOD, and DFX on Particulate-Induced 8-Oxo-dG Formation in Calf Thymus DNA incubation mixturea

8-oxo-dG/105 dGb

OFA and DNA OFA, DNA, and CAT OFA, DNA, and SOD OFA, DNA, and DFX CFA and DNA CFA, DNA, and CAT CFA, DNA, and SOD CFA, DNA, and DFX

26.49 ( 0.31 15.20 ( 0.20c 29.10 ( 0.21 12.80 ( 0.50c 9.75 ( 1.67 11.05 ( 0.05 9.70 ( 2.00 10.05 ( 0.75

a Particulates (1 mg/mL) were incubated with calf thymus DNA (0.5 mg/mL) for 15 min in the presence or absence of metal ion chelator DFX (0.2 mM) and antioxidant enzyme catalase (2000 units/mL) or SOD (30 units/mL) in phosphate buffer (pH 7.4), followed by enzymatic hydrolysis of DNA to nucleosides. The DNA hydrolysates were analyzed for dG and 8-oxo-dG levels by HPLC analysis as described in Materials and Methods. b The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. c Significant inhibition of 8-oxo-dG (p < 0.05).

SOD (Table 5). It is noteworthy that an overall trend of 8-oxo-dG formation in calf thymus DNA exposed to particulates mirrored dG hydroxylation results (see above). These results are in agreement with the notion that available or catalytically active metals associated with the particulates are critical in the pathway of particulate-induced dG hydroxylation and 8-oxo-dG formation in calf thymus DNA. Figure 6. Effect of OFA concentrations on the induction of 8-oxo-dG within calf thymus DNA. OFA at the indicated concentrations was incubated with DNA (0.5 mg/mL) in phosphate buffer (pH 7.4) for 15 min at room temperature under ambient air, followed by enzymatic hydrolysis of DNA to nucleosides. The DNA hydrolysates were analyzed for dG and 8-oxo-dG levels by HPLC analysis as described in Materials and Methods. The results are expressed as means ( the SD (n ) 3). Error bars represent the SD. The absence of an error bar indicates that the SD is contained within the symbol.

a HPLC eluant system containing 4 or 5% MeOH. The profiles compare the UV (lower trace) to EC detection (upper trace). It is interesting to note that the UV intensities of dC and T are relatively lower, exhibiting some sort of decomposition of pyrimidine nucleosides in particulate-treated DNA hydrolysate compared to the control. This trend is evident from the additional unknown minor UV peaks eluting before and after dC and after T as decomposition products (Figure 5B). Table 4 shows particulate-induced 8-oxo-dG formation within calf thymus DNA. A significant 4-fold increase in the level of 8-oxo-dG induction by OFA was noted over background levels in calf thymus DNA (p < 0.05). On the other hand, CFA had no effect (p > 0.05, compared to control DNA). Figure 6 illustrates the dose-dependent increase in the level of 8-oxo-dG formation in DNA exposed to OFA at concentrations of 0.1-1 mg/mL. As shown in Table 5, treatment of OFA (1 mg/mL) with metal ion chelator DFX (0.2 mM) before reaction with DNA or addition of a catalase (2000 units/mL) to the incubation mixture significantly suppressed 8-oxo-dG formation (p < 0.05). However, addition of SOD (30 units/ mL) to the incubation mixture slightly enhanced 8-oxodG formation in DNA, although the increase was not significant (p > 0.05). Contrary to these findings, the level of 8-oxo-dG never increased significantly above the control levels upon exposure of DNA to CFA (p > 0.05) or with particles treated with either DFX, catalase, or

Discussion An association between adverse health effects and respirable particulate matter at concentrations encountered in the urban environments is well recognized by epidemiologic findings (1-4). While precise mechanisms for these health effects remain unclear (5), there is a growing body of scientific evidence that provides some mechanistic plausibility for epidemiological health outcomes (30). In the study presented here, the oxidant generation capability of OFA and CFA in relation to particulate metal content and solubility was investigated in terms of their ability to induce dG hydroxylation and also 8-oxo-dG formation in calf thymus DNA following particulate exposure. The study examines 8-oxo-dG formation because this DNA base lesion is a widely accepted marker of oxidative damage and is usually used as a probe for the presence of •OH (22, 23, 31). In addition, other •OH-mediated DNA oxidation produces a wide spectrum of damage, including base lesions, oligonucleotide strand breaks, abasic sites, and DNA-protein cross-links (22, 23). Increasing interest has developed with regard to the role of metal ions in nucleic acid damage. Cadet et al. (23) have reviewed the complexity of the oxidation reactions that may be induced within the base moieties of DNA by metals. Most of the emphasis, however, has been placed on Fenton-driven Fe and Cu reactions (19). The transition metals Fe, Cu, Ni, V, Co, and Cr are capable of redox cycling electrons utilizing multiple valence states, which may initiate the production of ROS, such as O2-, H2O2, and highly reactive •OH, by reduction of molecular oxygen (8, 9). As a first approach to gain insight into the mode of action of particulate studied, we examined the efficiencies of different metal species in mediating dG hydroxylation to 8-oxo-dG formation at concentrations relevant to those detected in the fly ash particles.

Particulate-Induced Oxidative DNA Base Damage

For metals to facilitate the formation of ROS via the Fenton reaction, the metals must be in a free or catalytically active form (9). The soluble and reduced metal ions V(IV) or vanadyl and Fe(II) or ferrous exhibited relatively enhanced hydroxylation of dG to 8-oxo-dG compared to their counterpart V(V) or vanadate and Fe(III) or ferric ions. Unlike vanadyl or ferrous ions, Ni(II) is not easily oxidized to the +3 oxidation state under physiologic conditions (29). In the study presented here, significant induction of hydroxylation of dG to 8-oxo-dG by metal ions under ambient air was observed without exogenous H2O2 or metal reductants in the incubation mixture. However, these effects can be prevented in an oxygenfree atmosphere. Similar results were noted for vanadylor nickel(II)-mediated and Fe-containing mineral-mediated hydroxylation of dG (28, 29, 31). Our results indicate that molecular O2 is likely to be significantly involved in metal ion-catalyzed hydroxylation of dG and the underlying mechanism for •OH formation, in analogy with the Fenton reactions (8, 9). Preincubation of metals in 0.2 mM metal ion chelators, such as DFX, DTPA, and ferrozine, before reacting with dG, inhibited 8-oxo-dG formation (Table 1). These chelators are often used to determine whether metal ions are involved in the production of radicals. DFX and ferrozine are excellent chelators of Fe(III) or Fe(II) and have been used extensively in vitro to assess the role of iron toxicity (9). Many of the V-mediated biological effects have also been shown to be reduced or eliminated by DFX (6, 3236). DFX also significantly decreased the level of •OH generation from the vanadyl-mediated Fenton-like reaction (28). However, it should be mentioned that DFX scavenges •OH when present at high concentrations, such as 150 mM in cell culture (37). Since the concentration (0.2 mM) of DFX used in the current study was severalfold lower, it is highly unlikely that this chelator was functioning as a radical scavenger, though the system was acellular. Therefore, these results support the conclusion that soluble metals are critical in the generation of •OH. This radical is thought to be the major hydroxylating agent for dG and other nucleosides (38-40). Consistent with metal ion-mediated dG hydroxylation, OFA rich in soluble or ionizable V and Ni concentrations exhibited a dose-dependent increase in the level of hydroxylation of dG to 8-oxo-dG. Since Ni(II) is a poor hydroxylating agent compared to V and Fe cations (Table 1), it is reasonable to presume that much of the hydroxylating activity resulting from OFA comes from its soluble V and Fe content. Preincubation of OFA with metal ion chelators before addition to dG in buffer resulted in significant inhibition of 8-oxo-dG formation. We also demonstrated that addition of DMSO, a wellestablished •OH scavenger, to the particulate suspension containing dG dramatically suppressed 8-oxo-dG formation. These data convincingly support the conclusion that available or free metals associated with the particulate can generate highly reactive •OH, which in turn can react with dG to form 8-oxo-dG. At this stage, we do not know in what oxidation states these metals actually exist in particulate suspension. The soluble metals, for example, could exist in either V(IV) or V(V) and Fe(II) or Fe(III). Among air pollution particles, the elemental chemistry of coal fly ash particulate is much more complex and varies with coal type and source (20, 41). Conventional coal fly ash emitted into the atmosphere consists mainly

Chem. Res. Toxicol., Vol. 13, No. 10, 2000 1017

of insoluble Al, Si, Fe, and trace levels of other ionizable metals (20, 42). Despite the high sensitivity of the analytical method employed in the current study for 8-oxo-dG determinations, CFA containing high concentrations of insoluble Fe failed to induce significant hydroxylation of dG to 8-oxo-dG. These incubations, however, were carried out in the absence of either complexing agents or reductants, e.g., citrate or ascorbate. Studies have shown that addition of either complexing agents or reductants to the incubation mixture led to metal mobilization from crocidolite asbestos and several coal fly ash samples containing different amounts of otherwise nonavailable Fe (43, 44). The mobilized Fe from asbestos fibers has been implicated in Fe-catalyzed ROS induction, leading to oxidative damage to DNA (43). The reactivity of Fe and other metals is highly dependent upon the electronic environment. It has been demonstrated that •OH formation by vanadate via Fenton-like reactions, when performed in cacodylate buffer at physiological pH, sharply decreased the •OH yield (45). In contrast, •OH generation was highly favored, when the reaction was carried out in phosphate buffer (pH 7.4). Furthermore, Fe bound to low-molecular weight chelators such as citrate or EDTA has been shown to be redox active (46). Graf et al. (47) have shown that complexation of Fe by these chelators allows H2O, or other small molecules such as O2, to coordinate the metal. This enables the metal to reduce O2, generating highly reactive •OH. In contrast, when all the coordination sites of transition metals are bound by a chelator, as occurs with the metal ion chelators used in this study, the redox activity of these metals is drastically reduced. A significant increase in the level of 8-oxo-dG in the aerated aqueous dG solutions by a Fenton reagent in the presence of the reducing agent ascorbate has been explained in terms of competition between the reduction of the oxidizing guanylyl radical, an intermediate in 8-oxo-dG formation, and its reaction with molecular oxygen which is a rather slow process (22). The behavior of nucleosides cannot always be transposed to DNA, because of the role of stacking and other factors on the radical reactions of these compounds (22). However, in this study, particulate-induced dG hydroxylation findings paralleled the results from when calf thymus DNA was used as a substrate for 8-oxo-dG formation. Consistent with metal availability in particulates, OFA induced a dose-dependent increase in the extent of 8-oxo-dG formation in DNA. The treatment of the particles with DFX or addition of a catalase to the incubation mixture to decompose H2O2 attenuates the OFA effects. Addition of SOD slightly enhances 8-oxodG levels, indicating uptake of oxygen accompanied by production of O2- with concomitant dismutation to H2O2 and O2. Emphasis has been placed on the evaluation of mutagenic and carcinogenic effects of 8-oxo-dG (48, 49). However, it is likely that other oxidation products of both purine and pyrimidine bases are produced upon exposure of DNA to particulates. Indeed, it is apparent from Figure 5 that additional unknown minor UV peaks eluting before and after dC and after T suggest pyrimidine degradation following DNA exposure to particulates. These degradation products could be the result of deglycosylation of pyrimidines with the release of free bases as noted with dG by nickel compounds (29) or •OHmediated formation of hydroperoxides and corresponding hydroperoxyl pyrimidine radicals upon reaction with

1018

Chem. Res. Toxicol., Vol. 13, No. 10, 2000

oxygen. Some of these •OH-mediated degradation products of pyrimidine nucleosides have been characterized (22, 23). In line with dG hydroxylation data, CFA containing insoluble Fe did not induce significant 8-oxodG formation in isolated DNA. In conclusion, our results suggest that metal availability, but not the metal content in emission source particulates, is a critical factor in mediating molecular oxygen-dependent hydroxylation of dG to 8-oxo-dG and DNA base damage. To what extent the ROS-generating activity of these metal-containing air pollution particles and ambient dusts using model studies with dG and DNA models oxidative damage to DNA in vivo is currently under investigation.

Acknowledgment. We thank Drs. Stephen Nesnow and Michael Madden for their many helpful suggestions. A.K.P. was supported by the EPA/UNC Toxicology Research Program (CT902908). This paper has been reviewed by the National Health and Environmental Effects Research Laboratory, U.S. Environmental Protection Agency, and approved for publication. Approval does not signify that the contents necessarily reflect the views of the agency, nor does mention of trade names or commercial products constitute endorsement or recommendation for use.

References (1) Portney, P. R., and Mullahy, J. (1990) Urban air quality and chronic respiratory disease. Reg. Sci. Urban Econ. 20, 407-418. (2) Dockery, D. W., Pope, C. A., Xu, X., Spengler, J. D., Ware, J. W., Fae, M. E., Ferris, B. G., and Speizer, F. E. (1993) An association between air pollution and mortality in six U.S. cities. N. Engl. J. Med. 329, 1753-1759. (3) Schwartz, J. (1994) Air pollution and daily mortality: a review and meta analysis. Environ. Res. 64, 36-52. (4) Pope, C. A., Bates, D. V., and Raizenne, M. E. (1995) Health effects of particulate air pollution: Time for reassessment? Environ. Health Perspect. 103, 472-480. (5) Schlesinger, R. B. (1995) Toxicological evidence for health effects from inhaled particulate pollution: does it support the human experience? Inhal. Toxicol. 7, 99-109. (6) Ghio, A. J., Stonehuerner, J., Pritchard, R. J., Piantadosi, C. A., Quigley, D. R., Dreher, K. L., and Costa, D. L. (1996) Humic-like substances in air pollution particulates correlate with concentrations of transition metals and oxidant generation. Inhal. Toxicol. 8, 479-494. (7) Kadiiska, M. B., Mason, R. P., Dreher, K. L., Costa, D. L., and Ghio, A. J. (1997) In vivo evidence of free radical formation in the rat lung after exposure to an emission source air pollution particles. Chem. Res. Toxicol. 10, 1104-1108. (8) Meneghini, R. (1988) Genotoxicity of active oxygen species in mammalian cells. Mutat. Res. 195, 215-230. (9) Stohs, S. J., and Bagchi, D. (1995) Oxidative mechanisms in the toxicity of metal ions. Free Radical Biol. Med. 18, 321-336. (10) Pritchard, R. J., Ghio, A. J., Lehmann, J. R., Winsett, D. W., Tepper, J. S., Park, P., Gilmour, M. I., Dreher, K. L., and Costa, D. L. (1996) Oxidant generation and lung injury after particulate air pollutant exposure increase with the concentrations of the associated metals. Inhal. Toxicol. 8, 457-477. (11) Stock, P. (1960) On the relations between atmospheric pollution in urban and rural localities and mortality from cancer, bronchitis, pneumonia, with particular reference to 3,4-benzopyrene, beryllium, molybdenum, vanadium and arsenic. Br. J. Cancer 14, 397418. (12) Kraus, T., Raithel, H., and Schaller, K. H. (1989) Quantitation of manganese and vanadium levels in the lungs of normal and occupationally exposed persons. Zentralbl. Hyg. Umweltmed. 188, 108-126. (13) Leonard, A., and Gerber, G. B. (1994) Mutagenicity, carcinogenicity and teratogenicity of vanadium compounds. Mutat. Res. 317, 81-88. (14) Nechay, B. R., Nanninga, L. B., and Nechay, P. S. E. (1986) Vanadyl(IV) and vanadate(V) binding to selected endogenous phosphate, carboxyl, and amino ligands; calculations of cellular vanadium species distribution. Arch. Biochem. Biophys. 251, 128138.

Prahalad et al. (15) Kasprzak, K. S. (1991) The role of oxidative damage in metal carcinogenicity. Chem. Res. Toxicol. 4, 604-615. (16) Coogan, T. P., Latta, D. M., Snow, E. T., and Costa, M. (1989) Toxicity and carcinogenicity of nickel compounds. CRC Crit. Rev. Toxicol. 19, 341-384. (17) Kinlen, L. J., and Willows, A. N. (1988) Decline in the lung cancer hazard: A prospective study of the mortality of iron ore miners in Cumbria. Br. J. Ind. Med. 45, 219-224. (18) Boyd, J. T., Doll, R., Faulda, J. S., and Leiper, J. (1970) Cancer of the lung in iron ore miners. Br. J. Ind. Med. 27, 97-105. (19) Carmichael, A. J. (1990) Vanadyl-induced Fenton-like reaction in RNA an ESR and spin trapping study. FEBS Lett. 261, 165170. (20) Prahalad, A. K., Soukup, J. M., Inmon, J., Willis, R., Ghio, A. J., Becker, S., and Gallagher, J. E. (1999) Ambient air particles: Effects on cellular oxidant generation in relation to particulate elemental chemistry. Toxicol. Appl. Pharmacol. 158, 81-91. (21) Albrecht, D., and Jungi, T. (1993) Luminol-enhanced chemiluminescence induced in peripheral blood-derived human phagocytes: Obligatory requirement of myeloperoxidase exocytosis by monocytes. J. Leukocyte Biol. 54, 300-306. (22) Cadet, J. (1994) DNA damage caused by oxidation, deamination, ultraviolet radiation and photoexcited psoralens. In DNA Adducts: Identification and Biological Significance (Hemminki, K., Dipple, A., Shuker, D. E. G., Kadlubar, F. F., Segerback, D., and Bartsch, H., Eds.) IARC Scientific Publication 125, pp 245-276, International Agency for Research on Cancer, Lyon, France. (23) Cadet, J., Berger, M., Buchko, G. W., Incardona, M. F., Morin, B., Raoul, S., Ravanat, J. L., and Wagner, J. R. (1994) Metalcatalyzed oxidative degradation of DNA: Base damage and mechanistic aspects. In Trace Elements and Free Radicals in Oxidative Diseases (Favier, A. E., Neve, J., and Faure, P., Eds.) pp 20-36, American Oil Chemists’ Society, Champaign, IL. (24) Kasai, H., and Nishimura, S. (1984) Hydroxylation of deoxyguanosine at the C-8 position by ascorbic acid and other reducing agents. Nucleic Acids Res. 12, 2137-2145. (25) Fiala, E. S., Conaway, C. C., and Mathis, J. E. (1989) Oxidative DNA and RNA damage in the livers of Sprague-Dawley rats treated with the hepatocarcinogen 2-nitropropane. Cancer Res. 49, 5518-5522. (26) Marmur, J. (1961) A procedure for the isolation of deoxyribonucleic acid from micro-organisms. J. Mol. Biol. 3, 208-218. (27) Adachi, S., Zeisig, M., and Moller, L. (1995) Improvements in the analytical method for 8-hydroxydeoxyguanosine in nuclear DNA. Carcinogenesis 16, 253-258. (28) Shi, X., Jiang, H., Mao, Y., Ye, J., and Saffiotti, U. (1996) Vanadium(IV)-mediated free radical generation and related 2′deoxyguanosine hydroxylation and DNA damage. Toxicology 106, 27-38. (29) Kasprzak, K., and Hernandez, L. (1989) Enhancement of hydroxylation and deglycosylation of 2′-deoxyguanosine by carcinogenic nickel compounds. Cancer Res. 49, 5964-5968. (30) Schlesinger, R. B. (2000) Properties of ambient PM responsible for human health effects: Coherence between epidemiology and toxicology. Inhal. Toxicol. 12, 23-25. (31) Berger, M., de Hazen, M., Nejjari, A., Fournier, J., Guignard, J., Pezerat, H., and Cadet, J. (1993) Radical oxidation reactions of the purine moiety of 2′-deoxyribonucleosides and DNA by ironcontaining minerals. Carcinogenesis 14, 41-46. (32) Hansen, T., Aaseth, J., and Alexander, J. (1982) The effect of chelating agents on vanadium distribution in the rat body and on uptake by human erythrocytes. Arch. Toxicol. 50, 195-202. (33) Jones, M. M., and Bassinger, M. A. (1983) Chelates antidotes for sodium vanadate and vanadyl sulfate intoxication in mice. J. Toxicol. Environ. Health 12, 749-756. (34) Domingo, J. L., Llobet, J. M., and Corbella, J. (1985) Protection of mice against the lethal effects of sodium metavanadate: A quantitative comparison of a number of chelating agents. Toxicol. Lett. 26, 95-99. (35) Liochev, S. I., and Fridovich, I. (1990) Vanadate-stimulated oxidation of NAD(P)H in the presence of biological membrane and other sources of O2-. Arch. Biochem. Biophys. 279, 1-7. (36) Ghio, A. J., Meng, Z. H., Hatch, G. E., and Costa, D. L. (1997) Luminol-enhanced chemiluminescence after in vitro exposures of rat alveolar macrophages to oil fly ash is metal dependent. Inhal. Toxicol. 9, 255-271. (37) Morel, I., Cillard, J., Lescoat, G., Sergent, O., Pasdeloup, N., Ocaktan, A. Z., Abdallah, M. A., Brissot, P., and Cillard, P. (1992) Antioxidant and free radical scavenging activities of the iron chelators pyoverdin and hydroxypyrid-4-ones in iron-loaded hepatocyte cultures: comparison of their mechanism of protection with that of desferrioxamine. Free Radical Biol. Med. 13, 499-508.

Particulate-Induced Oxidative DNA Base Damage (38) Kasai, H., and Nishimura, S. (1984) DNA damage induced by asbestos in the presence of hydrogen peroxide. Gann 75, 841844. (39) Floyd, R. A., West, M. S., Eneff, K. L., Hogsett, W. E., and Tingey, D. T. (1988) Hydroxyl free radical mediated formation of 8-hydroxyguanosine in isolated DNA. Arch. Biochem. Biophys. 262, 266-272. (40) Loeb, L. A., James, E. A., Waltersdorp, A. M., and Klebanoff, S. J. (1988) Mutagenesis by the autoxidation of iron with isolated DNA. Proc. Natl. Acad. Sci. U.S.A. 85, 3918-3922. (41) Smith, K. L., Smoot, L. D., Fletcher, T. H., and Pugmire, R. J. (1994) The structure and reaction processes of coal. The Plenum Chemical Engineering Series, Plenum Press, New York. (42) Takahashi, S., Esaka, F., Sato, H., Kubota, Y., and Furuya, K. (1994) Concentrations of metal elements in mouse lung after intratracheal administration of coal fly ash. Inhal. Toxicol. 6, 6777. (43) Lund, L. G., and Aust, A. E. (1992) Iron mobilization from crocidolite asbestos greatly enhances crocidolite-dependent formation of DNA single-strand breaks in φX174 RFI DNA. Carcinogenesis 13, 637-642. (44) Smith, K. R., Veranth, J. M., Lighty, J. S., and Aust, A. E. (1998) Mobilization of iron from coal fly ash was dependent upon the

Chem. Res. Toxicol., Vol. 13, No. 10, 2000 1019

(45)

(46) (47)

(48)

(49)

particle size and the source of coal. Chem. Res. Toxicol. 11, 14941500. Ding, M., Gannett, P. M., Rojanasakul, Y., Liu, K., and Shi, K. (1994) One-electron reduction of vanadate by ascorbate and related free radical generation at physiological pH. J. Inorg. Biochem. 55, 101-112. Aust, A. E., and Eveleigh, J. F. (1999) Mechanism of DNA oxidation. Proc. Soc. Exp. Biol. Med. 222, 246-252. Graf, E., Mahoney, J. R., Bryant, R. G., and Eaton, J. W. (1984) Iron-catalyzed hydroxyl radical formation. J. Biol. Chem. 259, 3620-3624. Kuchino, Y., Mori, F., Kasai, H., Inoue, H., Iwai, S., Miura, K., Ohtsuka, E., and Nishimura, S. (1987) Misreading of DNA templates containing 8-hydroxydeoxyguanosine at the modified base and at adjacent residues. Nature 327, 77-79. Cheng, K. C., Cahill, D. S., Kasai, H., Nishimura, S., and Loeb, L. A. (1992) 8-Hydroxyguanine, an abundant form of oxidative DNA damage, causes GfT and AfC substitutions. J. Biol. Chem. 267, 166-172.

TX000110J