ETHANOL MODULATES RAT HEPATIC DNA REPAIR FUNCTIONS

Panida Navasumrit,1, Geoffrey P. Margison and Peter J. O'Connor,*

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


    ABSTRACT
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
— To further explore how ethanol may act at the DNA level, studies have been made of DNA repair mechanisms in male Wistar rats given ethanol either as an acute intragastric dose (5 g/kg) or continuously in a liquid diet (5% w/v) to provide 36% of the caloric intake. These treatments generate significant levels of free radicals with evidence of damage to DNA. The acute ethanol dose significantly inhibited O6-alkylguanine-DNA alkyltransferase (ATase) activity by 21–32% throughout the 24-h post-treatment period and this was confirmed by immunohistochemical detection of the ATase protein in hepatic nuclei. Twelve hours after the ethanol treatment, the activities of the DNA glycosylases, alkylpurine-DNA-N-glycosylase (APNG) and 8-oxoguanine-DNA glycosylase (OXOG glycosylase) were each increased by ~44%. In contrast, when given chronically via the liquid diet, ethanol initially had no effect on ATase activity, but after 4 weeks ATase activity was increased by 40%. Following ethanol withdrawal, ATase activity remained elevated for at least 12 h, but, by 24 h, the activity had fallen to the uninduced control level. DNA glycosylase activities were again affected differently. After 1 week of dietary ethanol exposure, there was no effect on APNG activity but it was inhibited by 19% at 4 weeks. OXOG glycosylase activity, on the other hand, was increased by 53% after 1 week, but decreased by 40% after 4 weeks. Although some of these changes in DNA repair capacity were relatively small, over time, their potential impact on the repair of endogenous or exogenous alkylation and/or oxidation damage in DNA would be substantial. These studies indicate possible mechanisms for the co-carcinogenic effects of ethanol.


    INTRODUCTION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Alcohol consumption is part of human lifestyle almost worldwide. Epidemiological studies show that the consumption of alcoholic beverages carries an associated increased risk of several cancers in humans and this increases with increasing levels of consumption (Boffetta and Garfinkel, 1990Go). Ethanol alone, however, has generally not been found to induce cancer in experimental animals (International Agency for Research on Cancer, 1988Go; Blot, 1992Go; Longnecker and Enger, 1996Go).

Ethanol oxidation gives rise to the generation of free radicals both in vitro and in vivo (Albano et al., 1991Go, 1996Go) 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., 2000Go). 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, 1989Go; Chinnasamy et al., 1996Go). 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., 1991Go; Chinnasamy et al., 1996Go). In another study, it was demonstrated that ATase gene induction is p53 gene-dependent: ATase activity was induced in mouse tissues following {gamma}-irradiation in p53 wild type mice, but not in p53 null animals (Rafferty et al., 1996Go). 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., 1993Go; Mitra and Kaina, 1993Go).

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.


    MATERIALS AND METHODS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Treatment of animals
Male outbred Wistar rats (130–150 g) from the Paterson Institute for Cancer Research (PICR) breeding unit were used under licence from the Home Office and with approval from the local Care and Use committee. For the acute ethanol treatment, ethanol 5 g/kg (25% w/v) was given by gavage and the controls received the same volume of vehicle. Chronic ethanol-treated animals were fed ethanol (5% w/v) in a liquid diet: this comprised 36% of total energy requirement, and the controls were pair-fed with an isocaloric liquid diet in which dextrin-maltose replaced the ethanol (Lieber and DeCarli, 1989Go). All animals were maintained on a 12-h light/12-h dark schedule and at designated times were killed by stunning and cervical dislocation.

Sera and antisera
Rat ATase antibodies were prepared at the Carcinogenesis Group, PICR. Goat anti-mouse IgG and goat anti-rabbit IgG horseradish peroxidase–antibody conjugate, swine anti-rabbit biotinylated immunoglobulins, avidin–biotinylated 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 Tris–HCl, 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, 1988Go). Protein concentrations were determined using bovine serum albumin (BSA) as a standard (Bradford, 1976Go).

Determination of APNG and OXOG glycosylase activities
For APNG assays, aliquots of tissue extracts containing 50–150 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 (20–30 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 sulphate–polyacrylamide 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., 1996Go).


    RESULTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Effects of ethanol treatment on ATase
Acute ethanol treatment. The effect of acute ethanol treatment on hepatic ATase activity and protein levels was studied at various times after ethanol administration as shown in Figure 1Go. ATase activity was decreased in ethanol-treated animals (Fig. 1AGo). The inhibition could be observed by 6 h and was significant at 12 and 18 h after treatment; thereafter the activity returned towards the control value. ATase protein levels were also assessed by Western immunoblotting (Fig. 1BGo). The immunoreactive band of ATase observed at ~22 kDa was more pronounced in protein extracts from the control group than those of ethanol-treated animals. The intensity of the ATase protein band decreased markedly from 6 to 18 h, but had increased again by 24 h after ethanol treatment, suggesting a relative delay in the effect on ATase protein (Fig. 1BGo).



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 1. Profile of hepatic O6-alkylguanine-DNA alkyltransferase (ATase) activity and immunoreactivity after acute ethanol treatment.

Wistar rats (~200 g) were treated with a single oral dose of ethanol (5 g/kg) and the same volume of water was given to the control group. ATase activity (fmol/mg protein; A) and immunoreactivity (B) were determined in the livers of either control (C) or ethanol-treated rats at 6, 12, 18, 24, 36 and 48 h after ethanol treatment. Each data point is the mean ± SEM from three to six animals. Protein from pooled samples of liver sonicates (three rats/group, 30 µg) was subjected to Western immunoblotting analysis as described in Materials and methods.

*Significantly different from the control value at P < 0.05.

 
Immunohistochemical staining also demonstrated that the intensity of ATase staining in both hepatic and some of the non-hepatic nuclei was reduced in the acutely ethanol-treated animals, as shown in Figure 2Go.



View larger version (74K):
[in this window]
[in a new window]
 
Fig. 2. Immunohistochemical detection of O6-alkylguanine-DNA alkyl-transferase (ATase) in the hepatic nuclei 12 h after acute ethanol treatment.

Negative control, section from an ethanol-treated animal (5 g/kg, intragastrically) stained with ATase-preadsorbed ATase antiserum (A); positive ATase-staining nuclei of ethanol-treated rat (B) or control rat (C). cv, central vein. Original magnification x200.

 
Chronic ethanol treatment. When animals were maintained on a liquid diet containing ethanol (5% w/v), this had no significant effect on hepatic ATase activity at 1, 2 and 3 weeks of treatment, but, after 4 weeks of feeding, it was induced significantly (Table 1Go). The increase was ~1.4-fold when the activity was expressed as a function of extract protein content.


View this table:
[in this window]
[in a new window]
 
Table 1. Time-course study on the effects of 5% (w/v) ethanol in a liquid diet on hepatic O6-alkylguanine-DNA alkyltransferase (ATase) activity
 
Over the treatment period the basal value of ATase activity also increased by ~1.7-fold in the pair-fed controls, possibly due to the effects of being maintained on a calorie-restricted diet. This compared to a ~2.6-fold increase in the ethanol-treated animals relative to protein concentration. (Table 1Go).

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. 3Go). Activity was elevated at time zero, increased further by 6 h after ethanol removal but then decreased thereafter (Fig. 3AGo). 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. 3BGo).



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 3. Effect of ethanol withdrawal on hepatic O6-alkylguanine-DNA alkyltransferase (ATase) activity and immunoreactivity in rats maintained for 4 weeks on ethanol in a liquid diet.

Animals were fed 5% (w/v) ethanol in a liquid diet or a control liquid diet for 4 weeks, then ethanol was removed from the diet. Hepatic ATase activity (fmol/mg protein; A) and immunoreactivity (B) were determined in the livers, either of pair-fed controls (C) or ethanol-treated rats at 0, 6, 12, 24 and 48 h. Each data point is the mean ± SEM from three to six animals. Protein from pooled samples of liver sonicates (three rats/group, 20 µg) was subjected to Western immunoblotting analysis as described in Materials and methods.

*Significantly different from control value at P < 0.05.

 
Effects of ethanol treatment on DNA glycosylase activities (APNG and OXOG glycosylase)
Acute ethanol treatment. In contrast to the effects on ATase, acute ethanol treatment increased the activity of both APNG and FPG. Twelve hours after acute ethanol treatment, the activities of APNG and OXOG glycosylase were increased by 42 and 44% respectively, compared to those of controls (Table 2Go).


View this table:
[in this window]
[in a new window]
 
Table 2. The effect of acute ethanol treatment on hepatic glycosylase activity
 
Chronic ethanol treatment. The effect of dietary ethanol on DNA glycosylase activity was also studied after 1 week and 4 weeks of feeding. OXOG glycosylase activity was induced significantly by ~50% in ethanol-treated animals, when compared to the pair-fed controls after 1 week of feeding, while APNG activity was not affected (Table 3Go). After 4 weeks of feeding, however, both were decreased significantly in ethanol-treated animals as shown in Table 3Go. In ethanol-fed animals, APNG activity was decreased by ~19% relative to the pair-fed control value, whilst the OXOG glycosylase activity was inhibited more strongly by ~40%.


View this table:
[in this window]
[in a new window]
 
Table 3. The effect of 5% (w/v) ethanol in a liquid diet on hepatic glycosylase activity
 

    DISCUSSION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
The current study indicated that ATase and the glycosylases (APNG and OXOG glycosylase) were affected by ethanol, resulting in differential elevation or inhibition of these DNA repair systems, depending on the circumstances of ethanol exposure. These differences indicate that expressions of these DNA repair proteins are controlled by different mechanisms, as will be discussed below.

ATase
Acute ethanol treatment inhibited ATase activity. This may be explained by the results of a previous study (Espina et al., 1988Go), 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 12–18 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., 1996Go; Major et al., 1997Go) 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., 2000Go). 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., 1985Go). ATase was also up-regulated by treatment of rat hepatoma H4IIE cells with H2O2 or by irradiation (Chan et al., 1992Go) 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., 1992Go). 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., 1993Go; Navasumrit et al., 2000Go). 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 {gamma}-irradiation to mice using systemically administered 114In (R. E. Wilson et al., 1994Go), 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., 1993Go).

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., 2000Go) 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, 1991Go; Laval, 1996Go).

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., 1993Go). Reactive radicals are also generated by acetaldehyde (Fridovich, 1989Go; Mira et al., 1995Go), 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., 1995Go; Nair et al., 1995Go).

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., 1998Go). In addition to the 3- and 7-alkylpurines, APNG also acts on stucturally unrelated oxidation products, e.g. ethenoadenine (Singer and Hang, 1997Go). 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., 2000Go). In addition, ethanol consumption can cause an iron overload in liver (Cederbaum, 1989Go), 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., 2001Go). 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, 1988Go). 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., 1993Go) 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, 1992Go), or as a urinary marker (Shuker et al., 1993Go), 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-{alpha} and nuclear factor-kB resulting in the generation of nitric oxide and the induction of apoptosis (French, 1996Go). It has been reported that nitric oxide inhibits OXOG glycosylase in E. coli and mammalian cells (Wink and Laval, 1994Go) as well as DNA ligase (Graziewicz et al., 1996Go). 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, 1994Go). 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., 1993Go). 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, 1991Go; Lefebvre et al., 1993Go) 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, 2000Go) 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, 1988Go).



View larger version (15K):
[in this window]
[in a new window]
 
Fig. 4. Comparison of the activities of O6-alkylguanine-DNA alkyltransferase (ATase) and the glycosylases [alkylpurine-DNA-N-glycosylase (APNG) and 8-oxoguanine-DNA glycosylase (OXOG glycosylase)] in rat liver 12 h after acute ethanol treatment.

Twelve hours after a single oral dose of ethanol (5 g/kg), ATase activity, APNG and OXOG glycosylase activities were determined. The specific activities were expressed as follows: ATase, fmol [3H-methyl] removed/mg protein; APNG and OXOG glycosylase, fmol [3H-methyl] released/mg protein; each bar represents the mean ± SEM from three to six animals. *Significantly different from the corresponding control value at P < 0.05.

 


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 5. Comparison of O6-alkylguanine-DNA alkyltransferase (ATase) activity and the glycosylase activities [alkylpurine-DNA-N-glycosylase (APNG) and 8-oxoguanine-DNA glycosylase (OXOG glycosylase)] in the liver of rats maintained for 1 or 4 weeks on 5% (w/v) ethanol in a liquid diet.

ATase, APNG and OXOG glycosylase activities in liver extracts of either pair-fed controls or ethanol-treated rats were analysed after 1 and 4 weeks feeding (A and B, respectively). Details are as described in the legend to Fig. 4Go. *Significantly different from the corresponding control value at P < 0.05.

 

    ACKNOWLEDGEMENTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
This work was supported by the Cancer Research Campaign (CRC), UK and by a grant to Panida Navasumrit from the government of Thailand.


    FOOTNOTES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
1 Present address: Chulabhorn Research Institute, Bangkok 10210, Thailand. Back

* Author to whom correspondence should be addressed. Back


    REFERENCES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Albano, E., Tomasi, A., Persson, J. O., Terelius, Y., Gatti, L. G., Ingelman-Sundberg, M. and Dianzani, M. U. (1991) Role of ethanol-inducible cytochrome P-450 (P450IIE1) in catalysing the free radical activation of aliphatic alcohols. Biochemical Pharmacology 41, 1895–1902.[ISI][Medline]

Albano, E., Clot, P., Morimoto, M., Tomasi, A., Ingelman-Sundberg, M. and French, S. W. (1996) Role of CYP2E1-dependent formation of hydroxylethyl free radical in the development of liver damage in rats intragastrically fed with ethanol. Hepatology 23, 155–163.[ISI][Medline]

Blot, W. J. (1992) Alcohol and cancer. Cancer Research 52, 2119s–2123s.[Abstract]

Boffetta, P. and Garfinkel, L. (1990) Alcohol drinking and mortality among men enrolled in an American Cancer Society prospective study. Epidemiology 1, 342–348.[Medline]

Bradford, M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72, 248–254.[ISI][Medline]

Cederbaum, A. I. (1989) Oxygen radical generation by microsomes: role of iron and implications for alcohol metabolism and toxicity. Free Radical Biology and Medicine7, 559–567.[ISI][Medline]

Chan, C. L., Wu, Z., Eastman, A. and Bresnick, E. (1992) Irradiation-induced expression of O6-methylguanine-DNA-methyltransferase in mammalian cells. Cancer Research 52, 1804–1809.[Abstract]

Chen, J. and Samson, L. (1991) Induction of S. cerevisae MAG 3-methyladenine DNA glycolsylase transcript levels in response to DNA damage. Nucleic Acids Research 19, 6427–6430.[Abstract]

Chinnasamy, N., Rafferty, J. A., Margison, G. P., O'Connor, P. J. and Elder, R. H. (1996) Induction of O6-alkylguanine-DNA-alkyltransferase in the hepatocytes of rats following treatment with 2-acetylaminofluorene. DNA and Cell Biology 16, 493–500.[ISI]

Espina, N., Lima, V., Lieber, C. S. and Garo, A. J. (1988) In vivo and in vitro inhibitory effect of ethanol and acetaldehyde on O6-methylguanine transferase. Carcinogenesis 9, 761–766.[Abstract]

Fraenkel-Conrat, H., and Singer, B. (1988) Nucleoside adducts are formed by co-operative reaction of acetaldehyde and alcohols: possible mechanism for the role of ethanol in carcinogenesis. Proceedings of the National Academy of Sciences of the USA 85, 3758–3761.[Abstract]

French, S. W. (1996) Ethanol and hepatocellular injury. Clinical Laboratory Medicine 16, 289–306.

Fridovich, I. (1989) Oxygen radicals from acetaldehyde. Free Radical Biology and Medicine 7, 557–558.[ISI][Medline]

Fritz, G. and Kaina, B. (1992) Stress factors affecting expression of O6-methylguanine-DNA methyltransferase mRNA in rat hepatoma cells. Biochimica et Biophysica Acta 1171, 35–40.[ISI][Medline]

Ghissassi, F., Barbin, A., Nair, J. and Bartsch, H. (1995) Formation of 1,N6-ethenoadenine and 3,N4-ethenocytosine by lipid peroxidation products and nucleic acid bases. Chemical Research in Toxicology 8, 278–283.[ISI][Medline]

Graziewicz, M., Wink, D. A. and Laval, F. (1996) Nitric oxide inhibits DNA ligase activity: potential mechanisms for NO-mediated DNA damage. Carcinogenesis 17, 2501–2505.[Abstract]

Grombacher, T. and Kaina, B. (1996) Isolation and analysis of inducibility of the rat N-methylpurine-DNA glycosylase promoter. DNA and Cell Biology 15, 581–588.[ISI][Medline]

Hall, J., Bresil, H., Donato, F., Wild, C. P., Loktionova, N. A., Kazanova, O. I., Komyakov, I. P., Lemekhov, V. G., Likhachev, A. J. and Montesano, R. (1993) Alkylation and oxidative-DNA damage repair activity in blood leukocytes of smokers and non-smokers. International Journal of Cancer 54, 728–733.[ISI]

Ingelman-Sundberg, M., Johansson, I., Yin, H., Terelius, Y., Eliasson, E., Clot, P. and Albano, E. (1993) Ethanol-inducible cytochrome P4502E1: genetic polymorphism, regulation, and possible role in the etiology of alcohol-induced liver disease. Alcohol 10, 447–452.[ISI][Medline]

International Agency for Research on Cancer (1988) Monographs on the Evaluation of the Carcinogenic Risks to Humans: Alcohol Drinking, vol. 44. International Agency for Research on Cancer, Lyon.

Laval, F. (1991) Increase of O6-methylguanine-DNA-methyltransferase and N3-methyladenine glycosylase RNA transcripts in rat hepatoma cells treated with DNA-damaging agents. Biochemical and Biophysical Research Communications 176, 1086–1092.[ISI][Medline]

Laval, J. (1996) Role of DNA repair enzymes in the cellular resistance to oxidative stress. Pathologie et Biologie, Paris 44, 14–24.

Lefebvre, P., Zak, P. and Laval, F. (1993) Induction of O6-methylguanine-DNA-methyl transferase and N3-methyladenine-DNA-glycosylase in human cells exposed to DNA-damaging agents. DNA and Cell Biology 12, 233–241.[ISI][Medline]

Lieber, C. S. and DeCarli, L. M. (1989) Review: Liquid diet technique of ethanol administration: 1989 update. Alcohol and Alcoholism 24, 197–211.[ISI][Medline]

Longnecker, M. P. and Enger, S. M. (1996) Epidemiological data on alcoholic beverage consumption and risk of cancer. Clinica Chimica Acta 246, 121–141.[ISI][Medline]

Major, G. N., Brady, M., Notarianni, G. B., Collier, J. D. and Douglas, M. S. (1997) Evidence for ubiquitin-mediated degradation of the DNA repair enzyme for O6-methylguanine in non-tumour derived human cell and tissue extracts. Biochemical Society Transactions 25, 359S.[Medline]

Margison, G. P., Butler, J. and Hoey, B. (1985) O6-Methylguanine methyltransferase activity is increased in rat tissues by ionising radiation. Carcinogenesis 6, 1695–1702.

Mira, L., Maia, L., Barreira, L. and Manso, C. F. (1995) Evidence for free radical generation due to NADH oxidation by aldehyde oxidase during ethanol metabolism. Archives of Biochemistry and Biophysics 318, 53–58.[ISI][Medline]

Mitra, S. and Kaina, B. (1993) Regulation of repair of alkylation damage in mammalian genomes. Progress in Nucleic Acid Research 44, 109–142.[ISI][Medline]

Morten, J. E. N. and Margison, G. P. (1988) Increased O6-alkylguanine-DNA-alkyltransferase activity in chinese hamster V79 cells following selection with chloroethylating agents. Carcinogenesis 9, 45–49.[Abstract]

Mustonen, R. and Hemminki, K. (1992) 7-Methylguanine levels in DNA of smoker's and non-smoker's total white blood cells, granulocytes and lymphocytes. Carcinogenesis 13, 1951–1955.[Abstract]

Nair, J., Barbin, A., Guichard, Y. and Bartsch, H. (1995) 1, N6-Ethenodeoxyadenosine and 3,N4-ethenodeoxycytidine in liver DNA from humans and untreated rodents detected by immunoaffinity/32P-postlabeling. Carcinogenesis 16, 613–617.[Abstract]

Navasumrit, P., Ward, T. H., Dodd, N. and O'Connor, P. J. (2000) Ethanol induced free radicals and hepatic DNA strand breaks are prevented in vivo by antioxidants: effects of acute and chronic ethanol exposure. Carcinogenesis 21, 93–99.[Abstract/Free Full Text]

Navasumrit, P., Ward, T. H., O'Connor, P. J., Nair, J., Frank, N. and Bartsch, H. (2001) Ethanol enhances the formation of endogenously and exogenously derived adducts in rat hepatic DNA. Mutation Research 479, 81–94.[ISI][Medline]

O'Connor, P. J. (1989) Towards a role for promutagenic lesions in carcinogenesis. In DNA Repair Mechanisms and their Biological Implications in Mammalian Cells, Lambert, M. W. and Laval, J. eds, pp. 61–71. Plenum Press, New York.

Pegg, A. E. (2000) Repair of O6-alkylguanine by alkyl transferases. Mutation Research 462, 83–100.[ISI][Medline]

Potter, P. M., Rafferty, J. A., Cawkwell, L., Wilkinson, M. C., Cooper, D. P., O'Connor, P. J. and Margison, G. P. (1991) Isolation and cDNA cloning of a rat O6-alkylguanine-DNA-alkyltransferase gene: molecular analysis of expression in rat liver. Carcinogenesis 12, 727–733.[Abstract]

Prieto Alamo, M. J., Jurado, J., Francastel, E. and Laval, F. (1998) Rat 7,8-dihydro-8-oxoguanine DNA glycolsylase: substrate specificity, kinetics and cleavage mechanism at an apurinic site. Nucleic Acids Research 26, 5199–5202.[Abstract/Free Full Text]

Rafferty, J. A., Clarke, A. R., Sellapan, D., Santibanez-Koref, M., Frayling, I. M. and Margison, G. P. (1996) Induction of murine O6-alkylguanine-DNA-akyltransferase in response to ionising radiation is p53 gene dose dependent. Oncogene 12, 693–697.[ISI][Medline]

Shuker, D. E. G., Prevost, V., Freisen, M. D., Lin, D. X., Oshima, H. and Bartsch, H. (1993) Urinary markers of exposure to endogenous and exogenous alkylating agents and precursors. Environmental Health Perspectives 99, 33–37.[ISI][Medline]

Singer, B. and Hang, B. (1997) What structural features determine repair enzyme specificity and mechanism in chemically modified DNA. Chemical Research in Toxicology 10, 713–732.[ISI][Medline]

Srivenugopal, K. S., Yuan, X. H., Friedman, H. S. and Ali-Osman, F. (1996) Ubiquination-dependent proteolysis of O6-methylguanine-DNA methyltransferase in human and murine tumour cells following inactivation with O6-benzylguanine or 1,3,-bis(2-chloroethyl)-1-nitrosourea. Biochemistry 35, 1328–1334.[ISI][Medline]

Wilson, D. M., Tentler, J. J., Carney, J. P., Wilson, T. M. and Kelley, M. R. (1994) Acute ethanol exposure suppresses the repair of O6-methylguanine DNA lesions in castrated adult male rats. Alcoholism: Clinical and Experimental Research 18, 1267–1271.[ISI][Medline]

Wilson, R. E., Hoey, B. and Margison, G. P. (1993) Ionizing radiation induces O6-alkylguanine-DNA-alkyltransferase mRNA and activity in mouse tissues. Carcinogenesis 14, 679–683.[Abstract]

Wilson, R. E., Hoyes, K. P., Morris, I. L., Hendry, J. H. and Margison, G. P. (1994) In vivo induction of O6-alkylguanine-DNA-alkyltransferase in response to indium-114m. Radiation Research 138, 26–33.[ISI][Medline]

Wink, D. A and Laval, J. (1994) The Fpg protein, a DNA repair enzyme, is inhibited by biomediator nitric oxide in vitro and in vivo. Carcinogenesis 15, 2125–2129.[Abstract]