One-electron oxidation is not a major route of metabolic activation and DNA binding for the carcinogen 7H-dibenzo[c,g]carbazole in vitro and in mouse liver and lung

Heather V. Dowty, Weiling Xue, Kathy LaDow, Glenn Talaska and David Warshawsky1

Department of Environmental Health, University of Cincinnati, Cincinnati, OH 45267-0056, USA


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
7H-Dibenzo[c,g]carbazole (DBC) is a potent multi-site, multi-species carcinogen present in a variety of complex mixtures derived from the incomplete combustion of organic matter. Like many carcinogens, DBC requires metabolic activation to an electrophilic species to exert its mutagenic and carcinogenic effects. One-electron oxidation, leading to the formation of radical cation intermediates, has been proposed as a mechanism of metabolic activation for DBC in vitro resulting in unstable DNA adducts. The purpose of this research was to determine whether one-electron oxidation is a mechanism of activation and DNA binding for DBC in vivo. Specific depurinating DBC–DNA adducts formed by one-electron oxidation were analyzed in mouse liver at 4 h following a single i.p. dose of 40 mg/kg of 11 µCi [14C]DBC. In addition to five previously published adduct standards, two newly identified adduct standards were characterized by mass spectrometry and NMR, namely DBC-6-N7-Ade and DBC-6-N1-Ade; however, neither was observed in mouse liver. Only the DBC-5-N7-Gua adduct was observed in mouse liver extracts at a level of 6.5 ± 1.8 adducts/106 nucleotides. In addition, the formation of AP sites and stable DBC–DNA adducts was analyzed in mouse liver and lung at 4, 12 and 24 h following a single i.p. dose of 0.4, 4 or 40 mg/kg DBC (n = 3/group). There was a distinct time– and dose–response of stable DBC–DNA adducts detected by 32P-post-labeling. There was not a clear dose–response for formation of AP sites; however, a significant increase over control levels was observed at the 4 and 40 mg/kg dose groups at 4 and 12 h post dosing, respectively. Quantitative comparisons indicate that the depurinating DBC-5-N7-Gua adduct constitutes ~0.4% of total adducts measured whereas the stable adducts detected by 32P-post-labeling constitute 99.6% of total adducts measured following a 40 mg/kg dose and a 4 h time-point. The data indicate that one-electron oxidation does occur in mouse liver in vivo. However, one-electron oxidation is a minor mechanism of activation in that the percentage of total adducts formed through this route constitutes a minor percentage of the total adducts formed.

Abbreviations: AP sites, apurinic sites; ARP, aldehyde reacting probe; DBC, 7H-dibenzo[c,g]carbazole; HRP, horseradish peroxidase; PAHs, polycyclic aromatic hydrocarbons.


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
One-electron oxidation has been proposed as a major mechanism of metabolic activation for a number of polycyclic aromatic hydrocarbons (PAHs) in initiating DNA damage, compared with other routes of activation, such as diol epoxide metabolites (1). This pathway results in the formation of radical cation intermediates which are electrophilic and bind to DNA (2). In addition, the DNA adducts formed by this route of activation are unstable leading to spontaneous depurination. While there is evidence to support a role for radical cations in vitro, relatively few studies have investigated the importance of one-electron oxidation leading to DNA damage in vivo (3,4).

7H-Dibenzo[c,g]carbazole (DBC) is a potent carcinogen that belongs to the class of N-heterocyclic polynuclear aromatic hydrocarbons formed during the incomplete combustion of organic matter. As such, it is a component of a variety of complex mixtures including tobacco smoke condensate, synthetic coal fuels, shale oil, polluted river sediments and coal tar (5). It is tumorigenic and carcinogenic in a number of different species including mouse, rat, hamster and dog (6). It is an extremely potent liver carcinogen producing mainly hepatocellular adenomas and carcinomas (7). Like many carcinogens, DBC requires metabolic activation to exert its mutagenic and carcinogenic effects (8). The metabolic profile of DBC is unique, resulting in the formation of primarily phenol intermediates (9). The 3-OH metabolite of DBC has been proposed as a proximate genotoxicant in the activation of DBC to bind to DNA (10). However, the low ionization potential, 7.3 eV (11) of DBC indicates that one-electron oxidation and the formation of radical cation intermediates is another possible mechanism of activation.

Chen et al. (12) demonstrated that DBC is activated by one-electron oxidation in vitro following microsomal or horseradish peroxidase (HRP) metabolism in the presence of DNA. In this system, DBC forms four DNA adducts which spontaneously depurinate. The adducts observed presumably arise from a radical cation with charge localization primarily at the 5-position of DBC, but also the 6-position, including DBC-5-N7-Gua, DBC-5-N7Ade, DBC-5-N3Ade and DBC-6-N7-Gua. Spontaneous depurination of these adducts would result in the formation of apurinic (AP) sites as the major type of DNA damage. While AP sites are the most frequent spontaneous lesion in DNA, they are also mutagenic in both prokaryotic and eukaryotic cells (13,14).

A number of stable DBC–DNA adducts are detected in the target tissues liver and lung by 32P-post-labeling (15). However, it is not known whether unstable DBC–DNA adducts formed by one-electron oxidation or the resulting AP sites also occur in these target tissues. In this study, the formation of specific depurinating DBC–DNA adducts formed by one-electron oxidation were analyzed in the target tissue liver. In addition, the resulting AP sites were compared with the level of stable DBC–DNA adducts detected in the target tissues liver and lung.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chemicals
7H-Dibenzo[c,g]carbazole was newly synthesized to >99% purity following the method of Katritzky and Wang (7). Aldehyde reacting probe (ARP) was purchased from Dojindo Inc. (Kumamoto, Japan). Deuterated-DMSO containing 0.1% tetramethylsilane (TMS) was purchased from Aldrich (Milwaukee, WI). All other reagents were purchased from either Fisher Scientific (Pittsburgh, PA) or Sigma Chemical Co. (St Louis, MO).

HRP activation
DBC was activated by HRP in the presence of calf thymus DNA. The reactions contained 20 mM Tris–HCl pH 7.4, 0.5 mg/ml calf thymus DNA, DBC (1, 10 or 100 µM) or DMSO, 5.0 PU/ml HRP and 0.5 mM H2O2. Reactions were initiated by the addition of H2O2 and incubated at 37°C for 1 h. The reactions were treated with 0.2 mg proteinase K for 1 h at 37°C. The DNA was then precipitated and resuspended in filtered ddH2O. The DNA was then analyzed for AP sites and stable adducts by ARP slot blot assay and nuclease P1 enhancement 32P-post-labeling assay, respectively.

AP site analysis
The aldehyde reacting probe slot blot assay was utilized to detect AP sites by following the methodology described by Nakamura et al. (16). AP site standards were generously provided by Jun Nakamura (Chapel Hill, NC). Briefly, 15 µg DNA (sample or standard) were incubated in 1 mM ARP in PBS for 10 min. DNA was precipitated with cold ethanol and resuspended in Tris–EDTA buffer at 3 µg/100 µl. DNA (100 µl) was heat denatured at 100°C for 5 min, quickly chilled on ice and mixed 1:1 with 2 M ammonium acetate. The samples were immediately applied to a supported nitrocellulose membrane presoaked in 20x SSC using a slot blot vacuum filter apparatus. The filter was rinsed in 5x SSC for 15 min at 37°C, dried and baked in a vacuum oven at 80°C for 1 h. The membrane was then blocked with 10 ml Tris–NaCl buffer containing BSA (20 mM Tris–HCl, 0.1 M NaCl, 1 mM EDTA, 0.5% casein, 0.25% BSA, 0.1% Tween-20, pH 7.5) at room temperature for 1 h. The membrane was then incubated in the same solution containing streptavidin-conjugated HRP at room temperature for 30 min. After rinsing the membrane with washing buffer (0.26 M NaCl, 1 mM EDTA, 20 mM Tris–HCl, 0.1% Tween-20, pH 7.5) for 15 min, enzymatic activity was measured using ECL detection reagents and visualized by incubating with film for 5–45 s. The resulting bands on film were quantitated by image analysis using a Chemimager 4000 (Alpha Innotech) and Chemimager software.

32P-post-labeling-nuclease P1
Duplicate DNA samples were hydrolyzed to 3'-monophosphate nucleotides by incubating with 0.25 U micrococcal nuclease and 2.5 µg spleen phosphodiesterase at 37°C for 6 h. Each mixture was incubated with 10 µl nuclease P1 cocktail (0.6 µg/µl nuclease P1, 0.054 mM ZnCl2, 0.075 M NaOAc) for 5 min. Each mixture was incubated with 5 µl post-labeling cocktail containing 1.5 µl PNK-buffer (200 mM Bicine, 100 mM DTT, 10 mM Spermidine, 100 mM MgCl2), 0.28 µl polynucleotide kinase (10.7 U/µl) and 20 µCi [32P]ATP in 10 mM Bicine pH 9.6 at 37°C for 40 min. Subsequently, each sample was treated with 3 µl apyrase (6.7 U/ml). Aliquots (18 µl) of each sample were applied to the origin 1.5 cm from the bottom of a PEI cellulose TLC plate.

Thin-layer chromatography
A 30 cm Whatman filter paper wick was attached to the top of the plate and developed in the D1 direction in 0.65 M sodium phosphate pH 6.0 overnight. The plates were rinsed, and each origin was excised and transferred to separate 10x10 cm PEI plates using a magnet-transfer technique. Plates were developed in the D3 direction with D3 solvent (3.6 M lithium formate, 8.5 M urea pH 3.5) followed by removal of the D1 transfer plate and two rinses in deionized water. The plates were dried, turned 90° in the D4 direction and developed with D4 solvent (0.8 M LiCl, 0.5 M Tris, 8.5 M urea, pH 8). The plates were then rinsed twice, dried and developed in the same direction with D5 solvent (1.5 M sodium phosphate monobasic, pH 6.0). The plates were dried, marked with fluorescent crayon and assembled into film cassettes. Plates were exposed to Fujichrome film overnight and developed.

DBC–DNA adduct standard synthesis and identification
Synthesis of adenine and guanine positional isomer adducts of DBC by iodine oxidation was conducted according to a method described by Chen et al. (12). Briefly, DBC (1 mmol) and deoxyguanosine (10 mmol) or adenine (5 mmol) were dissolved in 15 ml anhydrous dimethylformamide to which iodine (3 mmol/ml) was added dropwise with stirring. The mixture was heated to 50°C for 4 h under nitrogen, cooled and aqueous sodium thiosulfate added to remove the brown iodine color. The dried reaction mix was extracted three times with a mixture of ethanol/chloroform/acetone (2:1:1). Individual adducts were purified by HPLC using a YMC ODS-AQ 5 µm column (10x250) and a Waters HPLC system. The column was eluted with a gradient of 40% acetonitrile in water for 5 min followed by a linear gradient to 100% acetonitrile in 60 min. Purified adducts were analyzed by mass spectrometry and NMR. Mass spectrometry was conducted on a MALDI-TOF mass spectrometer (Micromass, TOFSE). Samples were mixed (1:1) with a saturated solution containing {alpha}-cyanol-4-OH cinnamic acid dissolved in 50% acetonitrile/water, 0.1% trifluoroacetic acid (0.5–1.0 µl). A nitrogen laser (337 nm {lambda}, 4 ns pulse width, 40 µJ/pulse) was used to desorb the sample ions. For NMR analysis, purified adducts were dissolved in deuterated DMSO:0.1% tetramethylsilane and analyzed with a 600 MHz NMR apparatus (Innova 600) at 25°C. Chemical shifts ({delta}) are reported relative to TMS (0 p.p.m.) and the coupling constants (J) are given in hertz.

DBC-6-N7Ade
1H-NMR: {delta} 5.914[s, 6-NH2(Ade)], 7.552 [dd, 1H, 11H], 7.619 [dd, 1H, 3H], 7.758 [dd, 1H, 12H], 7.831 [dd, 1H, 2H], 7.969 [d, 1H, 9H], 8.126 [d, 1H, 10H], 8.181 [s, 1H, 5H], 8.207 [d, 1H, 4H], 8.374 [s, 1H, 2H(Ade)], 8.723 [s, 1H, 8H(Ade)], 9.113 [d, 1H, 13H], 9.189 [d, 1H, 1H], 12.366 [s, 1H, 7H], J1,2 = 8.1 Hz, J2,3 = 7.6 Hz, J3,4 = 7.7 Hz, J8,9 = 8.7 Hz, J10,11 = 7.7 Hz, J11,12 = 7.3 Hz, J12,13 = 8.3 Hz. MS: M+ = 401.

DBC-6-N1-Ade
1H-NMR: {delta} 7.549 [dd, 1H, 11H], 7.623 [dd,1H, 3H], 7.695 [s, 1H, 8H(Ade)], 7.710 [d,1H, 8H], 7.750 [dd, 1H, 12H], 7.836 [dd, 1H, 2H], 7.953 [d,1H, 9H], 8.121 [d, 1H, 10H], 8.202 [d, 1H, 4H], 8.240 [s, 1H, 5H], 8.289 [s, 1H, 6-NH2(Ade)], 8.688 [s, 1H, 2H(Ade)], 9.104 [d, 1H, 13H], 9.179 [d, 1H, 1H], 12.214 [s, 1H, 7H], J1,2 = 7.8 Hz, J2,3 = 6.1 Hz, J3,4 = 7.5 Hz, J8,9 = 8.5 Hz, J10,11 = 7.4 Hz, J11,12 = 6.4 Hz, J12,13 = 7.8 Hz. MS: M+ = 401.

Animal treatment
Female Hsd:Icr(Br) mice were purchased from Harlan Sprague–Dawley (Indianapolis, IN) at 4–6 weeks of age and allowed to acclimate for 1 week prior to treatment. Animals were given food and water ad libitum and housed individually in plastic shoebox cages. Mice (n = 3) were given [14C]DBC in corn oil (11 µCi; 40 mg/kg) by i.p. administration. The animals were killed after 4 h and liver and lung tissue harvested and quickly frozen in liquid nitrogen. In a separate experiment, mice were given one bolus dose of DBC in corn oil (0.4, 4.0 or 40 mg/kg) or vehicle control by i.p. injection. Animals were killed by CO2 inhalation at 4, 12 or 24 h following treatment. Liver and lung were removed and quickly frozen in liquid nitrogen.

Soxhlet extraction
Liver tissue (1 g) from mice treated with [14C]DBC was pulverized under liquid nitrogen. The tissue was extracted with a soxhlet extraction apparatus for 48 h using ~20 ml of a mixture of chloroform:methanol (1:1). The extract was evaporated to dryness under vacuum. The residue was dissolved in DMSO by placing in a sonicator for 1–2 h. Another aliquot of tissue was homogenized in PBS and analyzed for DNA content by Hoechst 33258 (15).

HPLC analysis
The [14C]DBC liver extracts were assayed for the presence of adenine or guanine depurinating adducts by HPLC. The extracts were separated by a YMC ODS-AQ (6x250 mm) analytical column in an acetonitrile:water or methanol:water gradient at 1 ml/min. For the acetonitrile:water gradient, the column was eluted with 40% acetonitrile in water for 5 min followed by a linear gradient to 100% acetonitrile in 60 min. For the methanol:water gradient, the column was eluted with 40% methanol in water for 5 min followed by a convex (CV5) gradient to 100% methanol in 60 min. A Waters HPLC system equipped with a 616 pump, 996 photodiode array detector, U6K manual injector, 600S solvent controller and a Shimadzu RF-535 fluorescence detector was used to perform the analysis. An aliquot of each extract was separated and fractions were collected at 0.5 min intervals. Each fraction was analyzed by scintillation spectrometry. Due to low levels of the putative depurinating adducts in each of the liver extracts, the remaining extracts from each animal were pooled and injected on HPLC collecting 0.5 min fractions. These fractions were analyzed by fluorescence spectrometry. Fractions corresponding to the retention time and fluorescence spectra of known standards were reinjected on the acetonitrile:water gradient, collected and again analyzed by fluorescence spectrometry. The same fraction was then reinjected on a methanol:water gradient.

Fluorescence spectra
Fluorescence spectra were analyzed using an Hitachi F-4500 fluorescence spectrophotometer. Fluorescence excitation spectra were recorded at an emission wavelength of 400 nm and fluorescence emission spectra were recorded at an excitation wavelength of 278 nm. Synchronous scans of both standard and unknown peaks were achieved using a delta wavelength ({Delta}{lambda}) corresponding to their individual Stokes shift.

DNA isolation
Liver or lung tissue was homogenized in 1% SDS–100 mM EDTA to which proteinase K (0.2 mg) was added and incubated for 3 h at 55°C (post-labeling) or 12 h at 4°C (ARP assay). RNase A (0.24 mg) and RNase T1 (40 µg) were then added and the mixture incubated for 30 min at 37°C. The homogenate was cooled on ice followed by protein precipitation with 5 M ammonium acetate. The supernatant was transferred to a new tube and DNA precipitated with 1 vol 2-propanol. The DNA was then rinsed in 70% cold ethanol, dried and resuspended in ddH2O pH 7.0. The DNA concentration was adjusted to 0.5 µg/µl.

DNA hydrolysis
DNA (2 µg) was hydrolyzed to 3'-monophosphates using 2.5 µg calf spleen phosphodiesterase and 0.25 U micrococcal endonuclease. The hydrolysis mixture was incubated for 6 h at 37°C.

32P-post-labeling-carrier-free
The hydrolyzed DNA was labeled at the 5' position using 200 µCi [{gamma}-32P]ATP and 1.7 U polynucleotide kinase at pH 9.0 and incubated for 40 min at 37°C. Samples were applied to the origins of poly(ethyleneimine)-cellulose (PEI) plates. Chromatography was carried out as outlined above.


    Results
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 Abstract
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 Materials and methods
 Results
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 References
 
Comparison of stable adducts versus AP sites following one-electron oxidation activation of DBC in vitro
In this study, the formation of AP sites as measured by the ARP slot blot method was compared with the level of stable DNA adducts detected by 32P-post-labeling following activation of DBC by HRP in vitro. (Figure 1Go) There was a dose-dependent increase in both AP sites and stable DBC–DNA adducts, indicating that the response was a function of the amount of DBC binding to DNA. A marked difference was observed between the number of stable DNA adducts versus the number of AP sites in all treatment groups (Figure 1Go). A 6–18 fold excess of stable adducts compared with AP sites were noted between concentrations of 1–100 µM DBC. These results are clearly different than other reports which demonstrate that activation of DBC in vitro by HRP metabolism leads to a ratio of 2:1 depurinating adducts to stable adducts as measured by fluorescence line narrowing spectroscopy and 32P-post-labeling, respectively (12).



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Fig. 1. Comparison of AP sites and stable adducts in DNA following activation of DBC by HRP in vitro. DBC (0, 1, 10 or 100 µM) was activated by HRP in the presence of CT-DNA. AP sites were measured by the ARP slot blot assay. Stable DNA adducts were measured by 32P-post-labeling. Error bars reflect SEM.

 
DNA adduct standards
The synthesis of monosubstituted positional isomers of DBC and adenine or guanine was carried out by iodine oxidation. In addition to the four expected adenine adducts, two additional adducts were observed, namely DBC-6-N7-Ade and DBC-6-N1-Ade. In all, six adenine adducts were observed by HPLC separation and confirmed by retention times (Table IGo), MALDI-TOF mass spectrometry and 1H-NMR according to previously published data (12), including DBC-5-N7Ade, DBC-5-N3Ade, DBC-5-N1Ade, DBC-6-N3Ade. Several guanine peaks were observed by HPLC; however, only the DBC-5-N7-Gua adduct was confirmed by mass spectrometry and 1H-NMR.


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Table I. Retention times of DBC–DNA adduct standards following HPLC separation
 
DBC-6-N7-Ade
The spectra of this adduct suggest substitution between the 6 position of DBC and the N7 position of adenine (Figure 2BGo). The shielding effect of the NH2(Ade) resulting in a broad singlet at {delta}5.91 is indicative of substitution at the N7 position of adenine. The upfield shift of the 8H doublet coupled with an absence of shielding effect on the 4H doublet implies substitution at the 6 position.



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Fig. 2. (A) DBC; (B and C) NMR spectra.

 
DBC-6-N1-Ade
The characteristics of this adduct suggest substitution at the 6 position of DBC and the N1 position of adenine (Figure 2CGo). A downfield shift of the 5H proton, upfield shift of the 8H proton in conjunction with an absence of shielding effect on the 4H doublet, indicate substitution at the 6 position of DBC. A deshielding effect on the 2H(Ade) proton resulting in a downfield shift and a shielding effect on the 6-NH2(Ade) resulting in resonance to a downfield position is indicative of substitution at the N1 position of adenine.

Identification of DBC–DNA adducts in mouse liver
Liver extracts from mice (n = 3) treated with [14C]DBC (40 mg/kg; 11 µCi) were separated by analytical HPLC using an acetonitrile:water gradient. A minor peak was observed at 24.6 min that corresponds to the retention time (24.6 min) of the DBC-5-N7-Gua standard (Table IGo). This peak was observed in each of the treated animals (Figure 3AGo). When HPLC fractions of each liver extract were collected, radioactivity was associated with this peak (Figure 3BGo). The retention time of the unknown peak was the same as the standard, DBC-5-N7-Gua. Upon reinjection of the fraction and standard, separately, onto the acetonitrile:water gradient the same retention times were observed (Figure 4AGo). When this fraction was again injected on the methanol:water gradient, the retention time was identical to that of the standard, DBC-5-N7-Gua (Figure 4BGo). Comparison of fluorescence spectra of the DBC-5-N7-Gua standard and the unknown fraction gave identical results (Figure 5Go). Peaks corresponding to the other adduct standards were not observed in the liver extracts, but they may be below our limits of detection.



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Fig. 3. Identification of DBC-5-N7-Gua adduct in liver extracts of mice treated with [14C]DBC. Mice were treated i.p. with 40 mg/kg [14C]DBC (11 µCi) and killed at 4 h post-dosing. Livers were soxhlet-extracted with chloroform:methanol (1:1) for 48 h. Extracts were separated by HPLC using an acetonitrile:water gradient and monitored for (A) fluorescence intensity or (B) radioactivity. The arrow refers to the peak corresponding to the DBC-5-N7-Gua adduct.

 


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Fig. 4. Comparison of HPLC retention times. (a) Liver extract fraction; (b) DBC-5-N7-Gua adduct standard. (A) acetonitrile:water gradient; (B) methanol:water gradient. The arrow refers to the peak corresponding to the DBC-5-N7-Gua adduct standard.

 


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Fig. 5. Comparison of fluorescence spectra of liver extract fraction and DBC-5-N7-Gua adduct standard. (A) Fluorescence excitation spectrum monitored at [({lambda}Em) = 400 nm]; (B) fluorescence emission spectrum monitored at [({lambda}Ex) = 278 nm]; (C) synchronous fluorescence spectrum is monitored using a {Delta}{lambda} that equals the {lambda}Em of the peak with the shortest wavelength – {lambda}Ex of the peak with the longest wavelength.

 
Dose/time–response of stable DBC–DNA adducts in mouse liver and lung
Two-dimensional TLC autoradiograms (Figure 6Go) depict liver and lung DBC–DNA adducts following a single i.p. administration of 0 or 40 mg/kg. As expected, seven stable adducts were detected in mouse liver whereas three were observed in lung following i.p. administration. Adducts were numbered to correspond to previously published reports (15); however, an additional adduct positioned adjacent to adduct 1 was observed in the present study that had not been observed previously (15), designated here as adduct 1a. In addition, adduct 4 was not observed in this study. The major adducts in liver were adducts 6, 5 and 3. In lung, the major adducts were 2 and 3. DNA adducts were not observed in the control animals in either tissue.



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Fig. 6. Two-dimensional TLC autoradiograms of DBC–DNA adducts from mouse liver and lung. Mice were treated i.p. with 40 mg/kg DBC (B and D) or solvent control (A and C) and killed at 4 h post-dosing. Liver (C and D) and lung (A and B) DNA were analyzed by 32P-post-labeling. Chromatograms were exposed to film for 2–18 h.

 
There was a distinct dose–response between treatment groups. As dose increased by log intervals (i.e. 0.4, 4.0 and 40.0 mg/kg) the differences in DBC–DNA adduct levels, as measured by 32P-post-labeling, increased by approximately the same magnitude (Figures 7 and 8GoGo). In addition, a change in adduct level was observed as a function of time. At the highest dose in liver, the RAL index continued to increase for all adducts up to 24 h, whereas at the mid and low doses, RAL appears to be either declining or approaching a plateau by 24 h. At all doses in lung tissue, DNA adducts appear to be increasing up to 24 h.



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Fig. 7. Dose/time–response of mouse liver DBC–DNA adducts. Mice were treated i.p. with DBC (0, 0.4, 4 and 40 mg/kg) and killed at 4, 12 and 24 h post-dosing: (A) 0.4 mg/kg; (B) 4.0 mg/kg; (C) 40 mg/kg. DNA adducts in mouse lung were analyzed by 32P-post-labeling. Each data point represents the mean ± SEM of adducts 1a (closed circles), 1 (open circles), 2 (diamonds), 3 (open triangles), 5 (closed squares), 6 (open squares) and 7 (closed triangles).

 


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Fig. 8. Dose/time–response of mouse lung DBC–DNA adducts. Mice were treated i.p. with DBC (0, 0.4, 4 or 40 mg/kg) and killed at 4, 12 or 24 h post-dosing. DNA adducts in mouse lung were analyzed by 32P-post-labeling. (A) 0.4 mg/kg; (B) 4.0 mg/kg; (C) 40 mg/kg. Each data point represents the mean ± SEM of adducts 1 (circles), 2 (triangles), 3 (squares) and 6 (diamonds).

 
Dose/time–response of AP sites in mouse liver and lung following DBC administration in vivo
AP sites were observed in both liver and lung DNA (Figures 9 and 10GoGo). While there was not a clear dose–response in all treatment groups, the number of AP sites in liver DNA were elevated at both 4 and 12 h post dosing at the mid and high dose groups. A significant increase of AP sites in liver DNA was observed (P <= 0.05) following the 4.0 and 40.0 mg/kg DBC treatment groups at 4 and 12 h, respectively, post-dosing. A statistically significant increase in AP sites was not observed in the lung DNA of treated animals.



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Fig. 9. Apurinic sites in mouse liver DNA following DBC treatment. Mice were treated i.p. with DBC (0, 0.4, 4 or 40 mg/kg) and killed at 4, 12 or 24 h post-dosing. AP sites in mouse liver were analyzed by the ARP slot blot assay. Error bars represent SEM. *P <= 0.05 compared with vehicle control.

 


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Fig. 10. Apurinic sites in mouse lung DNA following DBC treatment. Mice were treated i.p. with DBC (0, 0.4, 4 or 40 mg/kg) and killed at 4, 12 or 24 h post-dosing. AP sites in mouse lung were analyzed by the ARP slot blot assay. Error bars represent SEM.

 

    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The interaction between metabolic activation and DNA damage is important in understanding basic mechanisms leading to mutagenesis and carcinogenesis by genotoxic compounds. This study has investigated whether one-electron oxidation is a mechanism of metabolism for the carcinogen DBC, resulting in the formation of unstable DNA adducts and AP sites in vitro and in vivo. This is a novel pathway of metabolic activation that has been largely supported by Cavalieri et al. (2,12). The significance of this pathway is presumed to be related to the levels of unstable DNA adducts produced in target tissues such as mouse skin (3,4). The resulting DNA damage from an unstable DNA adduct is an AP site. While AP sites are the most frequent spontaneous lesion in DNA, they are also mutagenic (13,14). The results of this work indicate that one-electron oxidation of DBC leading to the formation of unstable DBC–DNA adducts occurs in vitro and in mouse liver. This is quantitatively a minor (0.4%) pathway of activation and DNA binding.

In this study, a 6–18-fold excess of stable DNA adducts compared with AP sites were noted following activation of DBC with HRP in vitro. These results are in direct contrast with other reports that suggest a 2-fold excess of depurinating DBC–DNA adducts compared with stable DNA adducts in vitro (12). While the components and timing of the in vitro reactions were similar to other studies (12), it is not clear as to why such a quantitative discrepancy exists. Certainly different methodologies were used to quantitate the resulting DNA damage, i.e. measuring AP sites directly versus indirectly by quantitating specific depurinating DNA adducts. In addition, interlaboratory variability in absolute adduct levels exists with the 32P-post-labeling assay and may help to explain some of the quantitative differences observed between other studies (12) and the data presented here. Overall, the data presented here support the idea that unstable DNA adducts may be a minor mechanism of DNA damage by one-electron oxidation compared with stable DNA adducts in vitro.

Synthesis of DBC adenine adducts by iodine oxidation yielded the expected products namely, DBC-5-N7-Ade, DBC-5-N3-Ade, DBC-6-N3-Ade, DBC-5-N1-Ade and DBC-5-N7-Gua. Two additional adducts were also observed including DBC-6-N7-Ade and DBC-6-N1-Ade. Interestingly, the 5 or 6 positions of DBC were the only positions bound to adenine or guanine by iodine oxidation. This may indicate that charge density is localized at either of these positions.

Radical cation intermediates have been proposed as the activated metabolites formed by one-electron oxidation for a number of different PAHs, including DMBA, B[a]P, DB[a,l]P and DBC (4,3,12,17). Of these compounds only B[a]P has been examined both in vitro and in mouse skin, the major target organ for B[a]P-induced skin tumors (4). The major adducts observed in skin were the depurinating B[a]P–DNA adducts formed by a one-electron oxidation mechanism and not the diol-epoxide pathway of activation. These observations were intriguing because they contradicted a long held belief that the diol-epoxide pathway is the only route of activation for B[a]P. In addition, the implications of this work suggested that a very different type of DNA damage, namely AP sites, may contribute to the mutagenic and carcinogenic potency of B[a]P in mouse skin. The finding that DBC was activated by one-electron oxidation, not only by HRP but also by microsomal enzymes, suggested that this mechanism of activation may be occurring in the target tissues relevant to DBC.

Only the DBC-5-N7-Gua adduct was observed in mouse liver extracts at a level of 6.5 ± 1.9 depurinating adducts/106 nucleotides. It has been proposed that for many PAHs, the major depurinating DNA adducts may reflect mutation specificity. For example, the major B[a]P–guanyl adduct is presumed to be a major intermediate in production of G->T mutations. However, the major base-substitution mutation observed following DBC administration in mouse lung is an A->T transversion. This type of mutation does not support a role for the involvement of the unstable DBC-5-N7-Gua adduct observed in mouse liver. This does not exclude the possibility that mutations may arise from the resultant AP sites. For example, a wide array of mutations induced by AP sites are observed in eukaryotic cells, including base substitution, frameshift and deletion mutations (14). It is not known, however, whether the amount of damage produced by this mechanism would result in an increased frequency of mutations.

If one-electron oxidation participates in the activation of DBC in its target tissues, specific depurinating adducts and AP sites would be the primary DNA damage produced through this mechanism. AP sites are not detected by the method of 32P-post-labeling used for DBC–DNA adduct analysis. This suggests that while stable DBC–DNA adducts are detected in the target tissues liver, lung and skin, other types of DNA damage may be occurring that cannot be identified by 32P-post-labeling analysis.

The number of AP sites detected in mouse liver DNA following DBC administration was elevated above control levels for most treatment groups. However, in mouse lung, there was no significant difference between control and treated groups. This is unlike observations in cells following administration of MMS where a distinct dose–response was noted (16). Detecting AP sites in vivo, however, may be challenging since the repair of AP sites is thought to be a rapid process. If AP sites are being formed and also repaired at a rapid rate, then damage by this mechanism may not be measurable in vivo. However, significant differences were observed in mouse liver, indicating that the formation of excess AP sites is a plausible mechanism of DNA damage by DBC in vivo.

While the one-electron oxidation mechanism of metabolism was explored in this study it was also necessary to put this into the context of stable DBC–DNA adduct formation in vivo. DBC is carcinogenic in mouse liver following dermal administration and carcinogenic in lung and tumorigenic in liver following i.p. administration of 5–40 mg/kg. This carcinogenic potency is reflected in the levels of DNA adducts detected by 32P-post-labeling in these tissues. The structural characteristic of these adducts is still unknown; however, a 3-OH metabolite has been proposed as the proximate metabolite leading to stable DBC–DNA adducts (10).

DBC–DNA adducts were observed in liver and lung by 32P-post-labeling. As expected, adduct levels were more intense in the liver than in the lung. At the 4 h time-point and 40 mg/kg dose group, the relative adduct labeling index x106 of stable adducts in liver equaled 1843 ± 195. This level far exceeds the level of AP sites or the depurinating DBC-5-N7-Gua adduct observed at the same dose group and time-point. At the 40 mg/kg dose level and 4 h time-point, stable adducts detected by 32P-post-labeling contribute to 99.6% of total adducts measured, whereas depurinating adducts contribute only 0.4% (Figure 11Go).



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Fig. 11. Comparison of AP sites, stable adducts and the DBC-5-N7-Gua adduct in mouse liver. The data are expressed as either RALx106 or AP sites/106. Error bars represent SEM.

 
The time-course of adduct formation was much like that expected for a bolus dosing regimen. The increase in adduct formation is an integration of absorption, distribution and metabolic activation following i.p. administration. At the highest dose, the rate of change appears to steadily progress up to 24 h, whereas the low and mid doses appear to be reaching a maximum. Previous studies indicate that total DBC–DNA adducts peak around 24–72 h in liver following i.p. dosing, followed by a biphasic decline over a period of days persisting for up to several weeks (18).

The relative amounts of AP sites, DBC-5-N7-Gua adduct and stable DNA adducts detected by 32P-post-labeling were compared in mouse liver at the 40 mg/kg dose group and 4 h time point. Stable adducts outnumber AP sites or the DBC-5-N7-Gua adduct by roughly 300:1 (Figure 11Go). This observation clearly demonstrates that the stable DBC–DNA adducts detected by 32P-post-labeling are a major type of DNA damage. The unstable adduct was detected; however, it occurs at levels much lower than stable adducts. This indicates that one-electron oxidation leading to the formation of unstable DNA adducts and AP sites is not a major mechanism of DNA damage by DBC.


    Acknowledgments
 
We would like to thank Jun Nakamura for providing AP site standards; Mark Rance and Vajira Nanayakara for generating the NMR and MS data; Marlene Jaeger and Howard Shertzer for helpful discussions. This work was supported by NIH grants 2R01-ES04203-11, 5T32-ES07250, 1P30-ES06096-07 and P01 ES05652-08.


    Notes
 
1 To whom correspondence should be addressed Email: david.warshawsky{at}uc.edu Back


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received April 26, 1999; revised December 28, 1999; accepted December 30, 1999.





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