Cancer Research Campaign Carcinogenesis Group, Paterson Institute for Cancer Research, Christie Hospital (NHS) Trust, Manchester M20 4BX, UK
Received 4 December 2000; in revised form 21 February 2001; accepted 5 March 2001
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ABSTRACT |
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INTRODUCTION |
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Ethanol oxidation gives rise to the generation of free radicals both in vitro and in vivo (Albano et al., 1991, 1996
) and such events are associated with the induction of DNA damage, e.g. DNA strand breaks and the modification of DNA bases (Navasumrit et al., 2000
). Changes in DNA repair activities, in response to DNA damage, are well established. O6-Alkylguanine-DNA alkyltransferase (ATase) for example, the protein responsible for the repair of certain types of DNA alkylation damage, is induced in rats by a variety of genotoxic agents, including some hepatocarcinogens, such as N-nitrosodimethylamine, aflatoxin B1, and 2-acetylaminofluorene (O'Connor, 1989
; Chinnasamy et al., 1996
). Grombacher and Kaina (1996) indicated that human ATase mRNA expression was also increased by alkylating agents (e.g. N-methyl-N'-nitro-N-nitrosoguanidine and methyl methanesulphonate) and by ionizing radiation via the induction of the ATase promoter. Similarly, ATase mRNA expression in rat liver was increased in response to treatment with 2-acetylaminofluorene (Potter et al., 1991
; Chinnasamy et al., 1996
). In another study, it was demonstrated that ATase gene induction is p53 gene-dependent: ATase activity was induced in mouse tissues following
-irradiation in p53 wild type mice, but not in p53 null animals (Rafferty et al., 1996
). The DNA glycosylase, alkylpurine-DNA-N-glycosylase (APNG) is also inducible by a number of agents, including alkylating agents and X-rays (Lefebvre et al., 1993
; Mitra and Kaina, 1993
).
As a consequence of these and other observations, there is considerable interest in investigating DNA repair modulation as a possible risk factor in carcinogenesis. In the present study, we have used the rat as a model to examine the effects of both binge and chronic exposures to ethanol on DNA glycosylase and ATase activities.
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MATERIALS AND METHODS |
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Sera and antisera
Rat ATase antibodies were prepared at the Carcinogenesis Group, PICR. Goat anti-mouse IgG and goat anti-rabbit IgG horseradish peroxidaseantibody conjugate, swine anti-rabbit biotinylated immunoglobulins, avidinbiotinylated peroxidase complex and normal swine serum were purchased from Dako Immunoglobulins (High Wycombe, Bucks, UK).
Radiochemicals
[3H]N-Nitroso-N-methylurea (NMU; sp. act., 18.7 and 19.4 Ci/mol) was obtained from Amersham International plc and [3H]NMU methylated calf thymus DNA was prepared in the Carcinogenesis Group, PICR.
Chemicals
Ecoscint was from Mensura Tech. Ltd, Wigan, UK. Agarose was purchased from Scotlab, Coatbridge, UK, and the Western blotting detection kit and Rainbow protein molecular weight markers were obtained from Amersham International plc. All other chemicals were of analytical grade and were obtained from Sigma Chemical Company, Poole, Dorset, UK.
Quantification of ATase activity
Tissue (100 mg) in 1 ml of ice-cold buffer I (50 mM TrisHCl, pH 8.3; 1 mM EDTA; 3 mM DTT) containing 5 µg leupeptin was sonicated and 10 µl phenylmethylsulphonylfluoride (50 PMSF mM in ethanol) was then added to the sample. The protein extract was separated from the cell debris by centrifugation at 15 000 g at 4°C for 10 min and the supernatant was transferred to a fresh tube for assay, as previously described, using [3H]NMU (sp. act. 19.7 Ci/mmol) methylated calf thymus DNA as substrate (Morten and Margison, 1988). Protein concentrations were determined using bovine serum albumin (BSA) as a standard (Bradford, 1976
).
Determination of APNG and OXOG glycosylase activities
For APNG assays, aliquots of tissue extracts containing 50150 mg of protein and 10 µl of [3H]NMU-methylated DNA at a specific radioactivity of 18.7 Ci/mmol were made up to a final volume of 100 µl with glycosylase buffer (10 mM KCl, 70 mM HEPES pH 7.8, 1 mM dithiothreitol). After incubation at 37°C for 1 h, 30 µl of precipitant [2 M NaCl, 1 mg/ml BSA and 0.5 mg/ml calf thymus DNA] and 250 µl of absolute ethanol were added to the assay mixture, vortexed and then placed on dry ice for 20 min. The mixture was then centrifuged at 15 000 r.p.m., at 4°C for 15 min. A portion of the supernatant (300 µl) was pipetted into a scintillation tube and mixed with 3 ml of Ecoscint for counting of the radioactivity using an LKB scintillation counter.
For OXOG glycosylase assays, the procedure was the same as for the APNG assay, except for the substrate which was ring-opened [3H-methyl]7-methylguanine (sp. act. 3.95 Ci/mmol) and the incubation period was extended to 2 h. The glycosylase activity was expressed as fmol [3H-methyl] released, relative to the amount of protein or DNA.
Western immunoblotting analysis of ATase
Protein gel electrophoreses were performed on a Biorad mini gel system. Protein (2030 µg) of the pooled microsomal fraction (three rats/group), obtained from ethanol-treated rats or from pair-fed controls was resolved on 12% sodium dodecyl sulphatepolyacrylamide gel electrophoresis. Blots were probed overnight at 4°C with a dilution of the rabbit anti-rat ATase IgG fraction (2.5 µg/ml) in Tris-buffered saline (TBS), pH 7.5 containing 5% (w/v) non-fat milk powder. After immunoblotting, the membrane was incubated with goat-anti-rabbit horseradish peroxidase for 1 h and then extensively washed in TBS containing 0.1% (v/v) Tween-20 and once with TBS alone. The immunoreactive protein band was visualized by ECL detection and X-ray film exposure.
Immunohistochemical detection of ATase
Tissues were fixed in 4% formalin for 24 h prior to 70% ethanol fixation, processed and paraffin wax sections (3 µm) were cut onto 3-aminopropyltriethoxysilane-subbed slides, warmed at 37°C for 1 h and stored at room temperature. The sections were dewaxed, rehydrated with ethanol and then immunoreactivity was determined as described previously (Chinnasamy et al., 1996).
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RESULTS |
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As in the case of the acute ethanol treatment, ATase staining was confined to the nuclei. The increase in ATase activity in the livers of ethanol-fed animals observed after 4 weeks of feeding, however, was not sufficient to provide a clearly visible difference in the intensity of nuclear staining, and hence this is not shown.
The effects of ethanol withdrawal on ATase activity and ATase protein levels were determined at 0, 6, 12, 24 and 48 h after removal of ethanol following 4 weeks of feeding a liquid diet containing ethanol (5% w/v) (Fig. 3). Activity was elevated at time zero, increased further by 6 h after ethanol removal but then decreased thereafter (Fig. 3A
). A significant enhancement could still be observed 12 h after ethanol removal before it finally reached the control level by 24 h. These observations broadly corresponded with the changes in the intensity of the immunoreactive protein band (Fig. 3B
).
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DISCUSSION |
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ATase
Acute ethanol treatment inhibited ATase activity. This may be explained by the results of a previous study (Espina et al., 1988), which showed that acetaldehyde, an ethanol metabolite, inhibits ATase activity in vitro, possibly by direct interaction with the cysteine residue at the active site. D. M. Wilson et al. (1994), however, reported that acute ethanol treatment [at 3 h after intraperitoneal injection of 30% ethanol, 1 ml/80 g (or ~3 g/kg)] had no effect on ATase activity in intact male rats, but that in castrated animals it was decreased significantly, indicating a hormonal control of ATase activity.
The differing results obtained in intact animals probably reflect the dose of ethanol, which was lower and of a shorter duration than that used in the earlier study. In the present study, ATase activity was apparently inhibited at 6 h after a single dose of 5 g/kg, but this did not become significant until 1218 h, after which time the activity gradually returned to normal. The profile of ATase activity was paralleled by changes in the amount of ATase protein, as shown by Western blotting. This implies that the decrease in ATase activity after acute ethanol treatment was at least partly due to a transcriptional or translational down-regulation. This may be mediated by an irreversible binding of acetaldehyde to ATase which then results in a temporary depletion of the available pools of ATase protein, or an enhancement of the ubiquitin-mediated catabolism of ATase (Srivenugopal et al., 1996; Major et al., 1997
) which may explain the time delay between the activity and protein profile changes.
Hepatic ATase activity was increased when ethanol was given as a part of the diet, but a significant elevation of ATase activity in ethanol-treated rats was observed only after 4 weeks. Different regimes of ethanol treatment can thus lead to differing ATase responses, suggesting that ethanol can influence this repair system via various mechanisms. In the case of dietary ethanol, the increase of ATase activity is possibly due to the effect of ethanol-generated free radicals, which results in hepatic DNA strand breaks (Navasumrit et al., 2000). It has been shown that ionizing radiation can induce ATase activity in several rat tissues and it was suggested that radiation-induced strand breaks were responsible (Margison et al., 1985
). ATase was also up-regulated by treatment of rat hepatoma H4IIE cells with H2O2 or by irradiation (Chan et al., 1992
) and the effect of the latter was inhibited by dimethylsulphoxide which is a scavenger of OH, indicating that the OH may play a role in the radiation-induced increase in ATase activity (Chan et al., 1992
). In addition, Fritz and Kaina (1992) reported that the formation of DNA breaks mediated by oxidative stress is the ultimate signal for ATase expression, and this tallies with the generation of free radicals and hepatic DNA strand breaks throughout the 6-week study period of these experiments (Navarsumrit et al., 2000). These may arise via a mechanism that involves the induction of CYP 2E1 and its capacity to generate reactive oxygen species (Ingelman-Sundberg et al., 1993
; Navasumrit et al., 2000
). However, whilst the highest incidence of DNA strand breaks was observed at 1 week, an induction of ATase was not observed until after 4 weeks of feeding. The lack of ATase induction during the initial period of ethanol treatment is probably explained by the fact that ethanol (acting indirectly) produces much lower levels of DNA damage, compared to irradiation, which rapidly produces a substantial number of DNA breaks and so stimulates an early ATase response. When ethanol-induced DNA strand breaks are generated over a prolonged period, however, ATase can be induced as a cumulative, late stress response. This response is similar to that caused by continuous administration of low levels of
-irradiation to mice using systemically administered 114In (R. E. Wilson et al., 1994
), to generate low levels of DNA damage. In this case, ATase activity was not fully induced until 7 days, as compared with 48 h after single higher doses of external beam radiation (Wilson et al., 1993
).
The increase in ATase activity caused by chronic exposure to ethanol was maintained only up to 12 h following ethanol removal, after which time it decreased gradually, possibly reflecting the low level of strand breaks and their repair. The profile of ATase activity was in good agreement with the changes in the amount of ATase protein that was induced after 4 weeks of ethanol feeding and subsequently degraded following ethanol withdrawal. Taken together, the consequences of ethanol-generated free radicals and hence induced DNA strand breaks (Navasumrit et al., 2000) may result in the induction of ATase by a mechanism which possibly involves a transient increase in translation. The increase in ATase activity in untreated animals over the period of feeding was also of interest, as it occurred in concert with the increasing incidence of DNA strand breaks in controls. It may, thus, imply an age-dependent endogenous generation of oxidative damage and consequently an adaptive up-regulation response of the ATase protein.
APNG and OXOG glycosylases
As can be seen in the case of ATase, alterations of DNA glycosylase activity can also arise as a secondary response to cellular damage, i.e. exposure to DNA damaging agents, including carcinogens and UV light (Chen and Samson, 1991; Laval, 1996
).
In the case of ethanol, the DNA-damaging effects are mediated possibly via the induction of CYP 2E1 and its capacity to generate reactive oxygen species (Ingelman-Sundberg et al., 1993). Reactive radicals are also generated by acetaldehyde (Fridovich, 1989
; Mira et al., 1995
), so that the enhanced capacity for the oxidation of ethanol resulting from increased levels of CYP 2E1 would increase the production of acetaldehyde and reactive oxygen species. The latter have been shown to contribute to the formation of promutagenic oxidative DNA adducts, i.e. 8-hydroxyguanine mediated by OH, as well as the etheno adducts derived from lipid peroxidation (Ghissassi et al., 1995
; Nair et al., 1995
).
Apart from oxidative damage, CYP 2E1 also bioactivates certain N-nitroso compounds, some of which can be formed endogenously, and so the induction of CYP 2E1 could also lead to DNA alkylation damage arising via these endogenous sources. In the present work, the ethanol-containing liquid diet increased APNG and OXOG glycosylase activities in liver after 1 week of feeding. Formamidopyrimidine-DNA glycosylase activity is a function of the OXOG glycosylase protein, which also excises 8-oxo-guanine from 8-oxo-guanine:cytosine base pairs, and the cDNA encoding the protein has recently been isolated from rat liver (Prieto Alamo et al., 1998). In addition to the 3- and 7-alkylpurines, APNG also acts on stucturally unrelated oxidation products, e.g. ethenoadenine (Singer and Hang, 1997
). Thus, enhancement of both DNA repair activities might be part of an adaptive mechanism to protect cells from oxidative and alkylative DNA damage as a consequence of ethanol-induced CYP 2E1, which coincides temporally with the generation of reactive free radicals during dietary exposure to ethanol (Navasumrit et al., 2000
). In addition, ethanol consumption can cause an iron overload in liver (Cederbaum, 1989
), an event that could also promote the production of these oxidative adducts. For these reasons, therefore, the formation of ethanol-mediated 8-hydroxyguanine is a likely event and although this has not been reported so far, the involvement of ethanol in the generation of etheno adducts has already been demonstrated (Navasumrit et al., 2001
). Ethanol may also contribute directly towards the formation of DNA adducts as it may interact with its own metabolite, acetaldehyde, to form mixed acetal adducts covalently bound to the exocyclic amino groups of nucleosides (Fraenkel-Conrat and Singer, 1988
). In agreement with these observations, previous studies have reported that APNG and OXOG glycosylase activities were higher in peripheral blood leukocytes of smokers (Hall et al., 1993
) and this was correlated with significantly increased levels of 7-methylguanine in blood DNA and excretion of 3-methyladenine in urine in smokers, compared to non-smokers. In addition, 8-hydroxyguanine was detected in peripheral blood mononuclear cell DNA (Mustonen and Hemminki, 1992
), or as a urinary marker (Shuker et al., 1993
), after exposure to cigarette smoke.
In contrast to the effects of acute and short-term dietary treatments, APNG and OXOG glycosylase activities were decreased following 4 weeks of an ethanol liquid diet treatment. Chronic ethanol exposure causes hepatocellular injury via multifactorial processes, one of which involves the stimulation of tumour necrosis factor- and nuclear factor-kB resulting in the generation of nitric oxide and the induction of apoptosis (French, 1996
). It has been reported that nitric oxide inhibits OXOG glycosylase in E. coli and mammalian cells (Wink and Laval, 1994
) as well as DNA ligase (Graziewicz et al., 1996
). If this is the case, then ethanol-mediated OXOG glycosylase depression may be due, at least in part, to the effects of ethanol-induced nitric oxide (Wink and Laval, 1994
). It should be noted that both APNG and OXOG glycosylases showed the same trend of responses to ethanol exposure, suggesting that they may be co-regulated, whereas the changes in ATase activity occur in the opposite direction to those of APNG and OXOG glycosylases. Similarly also, a strong correlation between APNG and OXOG glycosylases and an inverse correlation between glycosylases and ATase was also found in smokers (Hall et al., 1993
). These observations strongly suggest that these two repair systems are controlled by different regulators. The alterations of APNG and OXOG glycosylase activities caused by DNA-damaging agents (Laval, 1991
; Lefebvre et al., 1993
) are likely to involve modulation of transcription, as has been demonstrated for APNG and ATase (see above). Although the promoter regions of these genes contain a number of transcription factor binding sites (e.g. Pegg, 2000
) the influence of ethanol and/or its metabolism on the expression and/or interaction of such factors has yet to be established.
In summary, we have shown that acute (binge) and chronic (controlled) ethanol consumption can substantially affect the levels of expression of several DNA repair proteins. This will impair the processing of DNA damage generated both by endogenous processes, that may themselves be affected by ethanol, and by exogenous factors, that may include ethanol. Such events are likely to be major contributing factors to the mechanisms of co-carcinogenesis by this widely consumed agent, which is recognized as a human carcinogen (International Agency for Research on Cancer, 1988).
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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* Author to whom correspondence should be addressed.
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