CHRONIC CONSUMPTION OF ETHANOL LEADS TO SUBSTANTIAL CELL DAMAGE IN CULTURED RAT ASTROCYTES IN CONDITIONS PROMOTING ACETALDEHYDE ACCUMULATION

N. SIGNORINI-ALLIBE, B. GONTHIER, F. LAMARCHE, H. EYSSERIC and L. BARRET*

Laboratoire ORSOX (Oligoéléments et Résistance au Stress Oxydant induit par les Xénobiotiques) UMR UJF/CEA-LCR CEA 8M, Université Joseph Fourier, Domaine de la Merci, 38706 La Tronche Cedex, France

* Author to whom correspondence should be addressed at: Fédération de Toxicologie Clinique et Biologique, CHU de Grenoble, 38043 Grenoble Cedex, France. Tel.: (33) 4 76 765783; Fax: (33) 4 76 765177; E-mail: Luc.Barret{at}ujf-grenoble.fr

(Received 7 October 2003; first review notified 15 December 2003; in revised form 11 February 2004; accepted 1 September 2004; Advance Access publication 17 March 2005)


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Aims: This study aimed at comparing the cerebral cytotoxicity of ethanol and its main metabolite acetaldehyde after acute or chronic exposures of rat astrocytes in primary culture. Methods: Cytotoxicity was evaluated on the cell reduction of viability (MTT reduction test) and on the characterization of DNA damage by single cell gel electrophoresis (or comet assay). Results: Changes in astrocyte survival and in DNA integrity only occurred when the astrocytes were chronically exposed to ethanol (20 mM; 3, 6 or 9 days). On the other hand, viability and DNA integrity were deeply affected by acute exposure to acetaldehyde. Both effects were dependent on the concentration of acetaldehyde. The cytotoxic effect of acetaldehyde was also indirectly evaluated after modifications of the normal ethanol metabolism by the use of different inducers or inhibitors. In presence of ethanol, the concomitant induction of catalase (i.e. by glucose oxidase) and inhibition of aldehyde dehydrogenase (i.e. by methylene blue) led to acetaldehyde accumulation within cells. It was followed by both a reduction in viability and a substantial increase in DNA strand breaks. Conclusions: These data were thus consistent with a possible predominant role of acetaldehyde during brain ethanol metabolism. On the other hand, the effects observed after AMT could also suggest a possible direct ethanol effect and a role for free radical attacks. These data were thus consistent with a possible predominant role of acetaldehyde during brain ethanol metabolism. On the other hand, the effects observed after AMT could also suggest a possible direct ethanol effect and a role for free radical attacks.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The adverse effects of alcohol on different functions of the central nervous system have been well documented. However, the mechanism underlying these effects has yet to be thoroughly investigated. Several mechanisms have been proposed involving for example the enzyme-catalyzed oxidation of ethanol to acetaldehyde. Acetaldehyde is a highly reactive metabolite that may induce cytotoxic conditions by binding tissue macromolecules and disrupting cellular functions (Lieber, 1997Go). In brain, ethanol can be metabolized to acetaldehyde via two pathways: either catalase (Gill et al., 1992Go; Aspberg et al., 1993Go) or cytochrome P450 (Montoliu et al., 1995Go; Upadhya et al., 2000Go). Significant acetaldehyde production is characteristic of brain homogenates (Zimatkin et al., 1998Go) and in primary culture of astrocytes after acute and chronic exposure to ethanol (Eysseric et al., 1997Go). Acetaldehyde was further metabolized to acetic acid by the NAD-dependent enzyme aldehyde dehydrogenase (ALDH) (Lieber, 1982Go).

This study's aim was to characterize the ethanol and acetaldehyde impact on astrocytes by measuring cell cytotoxicity and DNA alterations taken as an early marker of ethanol toxicity. The effects of acetaldehyde were evaluated using two modes of exposure: either a direct cell exposure in the culture medium (at relatively low doses) or an indirect stress after modifications of ethanol metabolism, leading to an intracellular accumulation of acetaldehyde. Results were compared to those obtained after direct ethanol exposure.

Astrocytes were chosen as a model for cerebral intoxication because they are a specific target with considerable potential consequences. Indeed, the brain cell population is largely made of astrocytes. They also play an important role in the developmental guidance of migrating neurons, the regulation of neurotransmitters and ion levels, the nutrition of neurons, and the production of neurotrophic factors (Kimelberg and Norenberg, 1989Go). Astrocytes may also be a major site for the detoxification or the bioactivation of neurotoxins (Di Monte et al., 1996Go).

The advantage of primary cultures lies in their closer resemblance to cells found in vivo. The drawbacks of this method are their limited survival time and the continual need to prepare new cells in culture. This can explain why very few reports are found in the literature on this subject. In particular, no data are available concerning the genotoxicity of ethanol on brain cells at such low doses and such short exposure times in chronic conditions (ethanol 20 mM for 3, 6 or 9 days). Indeed, Russo et al. (2001)Go used ethanol concentrations of 50, 100 and 200 mM for 10 days, whereas Renis et al. (1996)Go fed rats with a diet containing 5% ethanol for 40 days. To better understand mechanisms of ethanol toxicity on astrocytes, both cell viability and nuclear DNA damage were thus investigated by inducing or inhibiting different pathways of ethanol metabolism.


    METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Culture of primary astrocytes
Pregnant Sprague–Dawley rats were obtained from Iffa-Credo (L'Arbresle, France). Animal care and use as well as all procedures involving animals were carried out according to national regulations.

The primary astrocyte cultures were prepared aseptically from cerebral hemispheres of 1- or 2-day-old pups, according to previously described methods (Booher and Sensenbrenner, 1972Go), with a few modifications (Gonthier et al., 1997Go).

The dissociated cells were plated in 35-mm diameter Petri dishes or 25-cm2 plastic tissue culture flasks at a density of 6.104 viable cells per cm2 in the usual D-MEM medium containing 10% FCS. The cultures were maintained at 37°C in a 5% CO2 humidified atmosphere. The medium was changed 24 h after seeding and then every 3 days. The cultures reached confluency after 7 days in vitro (DIV). On day 14, the monolayers were composed of 95% astrocytes as demonstrated by positive immunostaining with antiserum to {alpha}-GFAP, an astrocyte marker (Vijayan et al., 1993Go).

Conditions for ethanol exposure
All experiments were carried out in culture medium between 11 and 15 days in vitro and in an air/CO2 incubator. Each type of experiment was conducted simultaneously in vitro so that dates could be compared and statistical analysis carried out properly. Cells were acutely or chronically exposed to ethanol.

Acute alcohol exposure in D-MEM medium containing ethanol at a final concentration of 100 mM lasted for 3 h.

In conditions of chronic ethanol exposure, the cells were incubated in a culture medium containing 20 mM ethanol for 3, 6 or 9 days. To avoid ethanol evaporation, we used a previously described compensating system which ensured a constant concentration of alcohol in the culture medium for 3 days (Eysseric et al., 1997Go).

Conditions for acetaldehyde exposure
In these experiments, cells were exposed to acetaldehyde in tissue culture flasks. Acute acetaldehyde exposure was carried out by adding 0.25, 0.50 and 1 mM of acetaldehyde in D-MEM medium for 3 h. In order to avoid evaporation, because of the high volatility of acetaldehyde at temperatures above 20°C, the culture flasks were completely filled with the medium and hermetically capped. Despite these unfavourable conditions, no impact on the cell viability was observed for exposures lasting only 3 h.

Enzymatic catalysis
To specify the role of the different metabolic pathways implicated in the toxicity of ethanol and the particular role of acetaldehyde, the action of various inducers and inhibitors of enzymes implicated in the cerebral metabolism of ethanol were tested. All reagents were purchased from Sigma (St Quentin-Fallavier, France).

3-amino-1,2,4-triazole (AMT), a well-known catalase inhibitor, was added to the culture medium 10 h before ethanol at a final concentration of 10 mM (Gill et al., 1992Go). In order to generate hydrogen peroxide (the only catalase cofactor), glucose oxidase (GO, EC 1.1.3.4) was added, at a final concentration of 6 mU/ml for acute exposures and 3 mU/ml for chronic exposures, 1 h before ethanol in the culture medium containing 33 mM glucose (Pinteaux et al., 1996Go). Methylene blue, which acts as an aldehyde dehydrogenase inhibitor, was added 10 h before ethanol at a final concentration of 30 µM for acute exposures and 1 µM for chronic exposures (Helander et al., 1993Go).

Determination of cell viability
The viability of the astrocytes after different stresses was determined using the MTT reduction test (Iselt et al., 1989Go). For each condition of exposure, at least five dishes or flasks were tested.

Evaluation of DNA damage
Single cell gel electrophoresis. DNA damage was evaluated using the comet assay (McKelvey-Martin et al., 1993Go) undertaken immediately after the stress in order to prevent repair mechanisms from taking action. Therefore the effects on DNA observed in these conditions reflect the initial DNA damage in term of strand breaks and oxidatively damaged bases generating alkali-labile sites in DNA.

The procedure used was a modification of the protocol described by Singh et al. (1988)Go. Frosted microscope slides were first covered with 150 µl of 1% normal agarose in Ca2+- and Mg2+-free phosphate-buffered saline (PBS) and immediately covered with a 22 x 50 mm coverslip and kept at room temperature to allow the agarose to solidify. The coverslip was then gently slid off. About 20 000 cells were suspended in 80 µl of 0.8% low-melting point agarose in PBS kept at 37°C and transferred onto the first agarose layer. After covering with a coverslip, the slides were left on ice for 5 min. The coverslips were then removed and the slides were placed in freshly prepared lysing solution at 4°C for 1 h in the dark (2.5 M NaCl, 100 mM Na2EDTA, 10 mM Tris, 1% sodium sarcosinate). We added 10% DMSO and 1% Triton X-100 to this lysing solution just before use. After lysis the slides were gently transferred to a horizontal gel electrophoresis tank filled with freshly prepared electrophoresis solution (300 mM NaOH, 1 mM Na2EDTA, pH > 13) at room temperature in the dark. The DNA was allowed to unwind for 20 min and electrophoresis was carried out by adjusting the voltage to 25 V and the current to 300 mA for 15 min. After electrophoresis the slides were washed gently to remove alkali and detergents that would interfere with ethidium bromide staining, using neutralization buffer (0.4 M Tris–HCl, pH 7.4) three times for 5 min. After neutralization, the slides were stained with 50 µl of 3.3 µg/ml ethidium bromide in distilled water and covered with a coverslip. The slides were placed in a humidified air-tight container to prevent drying of the gel, until analysis. Three slides were prepared per assay and 50 nuclei were counted per slide.

Slide analysis. Slides were examined using an epifluorescence microscope, Zeiss Axioskop 20 (Carl Zeiss, Microscopre Division, Oberkochen, Germany), equipped with a short arc mercury lamp HBO (50 W, 516–560 nm, Zeiss), and filters 5 and 15 (Zeiss) at 20x magnification. Fifty randomly selected comets on each triplicate slide were scored with a Pulmix TM 765 camera (Kinetic Imaging, Liverpool, UK) linked to an image analysis system Komet 3.0 (Kinetic Imaging). This software defined different parameters for image processing. Among these parameters, we chose the percentage of DNA in the tail for the evaluation of DNA damage. The percentage of DNA in the tail is linearly related to DNA break frequency (Wozniak and Blasiak, 2003Go).

Statistics
Three independent experimental series were conducted for each condition of exposure, unless otherwise indicated. Controls corresponded to astrocytes exposed to neither ethanol nor acetaldehyde and grown in culture under the same conditions as in the experimental series.

Results were expressed as mean ± SEM. The statistical significance of the results was tested by an analysis of variance for the factorial model completed by the Fisher PLSD post hoc test to compare the mean values between the different experimental series and by the Mann–Whitney test when appropriate. The Kolgomorov–Smirnov test was employed to compare the distribution of the percentages of DNA in the tail.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Acute exposure to ethanol ± inhibitors and/or inducers of ethanol metabolism
The MTT assay showed that cell viability was not affected after ethanol exposure whatever the concentration tested (20, 50 or 100 mM for 3 h). We also observed an absence of significant DNA strand break formation as attested by single cell gel electrophoresis (data not shown).

The effects on cell viability and DNA damage induced by ethanol in presence of the inducers and/or inhibitors tested were strictly compared to the effects of these compounds on cells not exposed to ethanol. Under such conditions, no modification was observed after acute ethanol exposure (data not shown).

Acute exposure to acetaldehyde
The exposures to acetaldehyde were conducted in culture flasks completely filled with the culture medium and hermetically capped in order to minimize acetaldehyde evaporation. Despite these conditions, no significant impact on cell viability was observed in the absence of acetaldehyde.

The cytotoxic profile of acetaldehyde was similar to that of ethanol (Fig. 1A). Unlike acute alcohol exposure, acetaldehyde exposure (0.25, 0.5 or 1 mM for 3 h) induced DNA damage. Indeed, the results of the comet assay, shown in Fig. 1(B), showed a dose-dependent increase in the frequency of single- and double-DNA strand breaks and alkali-labile sites (P < 0.001, Anova test). The Fisher PLSD test showed values different from controls, especially for the highest doses of 0.5 and 1 mM (P < 0.0001).



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Fig. 1. (A) Astrocyte variation in viability after exposure for 3 h to various concentrations of acetaldehyde. Results are the mean of three independent experimental series and are expressed as mean ± SEM (n = 9). No significant difference was observed using the Fisher PLSD test. Control without acetaldehyde represents 100% viability. Experimental conditions are detailed in the Methods. (B) Level of DNA damage in rat astrocytes exposed for 3 h to various concentrations of acetaldehyde and analysed using the comet assay. DNA damage is expressed as percentage of DNA in the tail. The figure shows mean results of three independent experiments. Fifty cells were randomly examined in quintuplicate for each condition and results are expressed as mean ± SEM (*P < 0.05, ***P < 0.0001 when groups exposed to acetaldehyde are compared to control: Fisher PLSD test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 
This is confirmed in Fig. 2, which shows progressing distribution of the percentage of tail DNA depending on acetaldehyde concentration. Indeed, the most abundant fraction of the cells in control had tail DNA between 0% and 10% (Fig. 2A), whereas the most abundant fraction of the cells exposed to 1 mM acetaldehyde had tail DNA between 10% and 30% (Fig. 2D) (P < 0.0001, control versus 1 mM; Kolgomorov–Smirnov test).



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Fig. 2. Histograms of the distribution of comet tail DNA of astrocytes incubated for 3 h with various concentrations of acetaldehyde. (A) Control; (B) acetaldehyde 0.25 mM (B vs A, P < 0.01, Kolgomorov–Smirnov test); (C) acetaldehyde 0.5 mM (C vs A, P < 0.0001, C vs B, NS, Kolgomorov–Smirnov test); (D) acetaldehyde 1 mM (D vs A and D vs B, P < 0.0001, D vs C, P < 0.05, Kolgomorov–Smirnov test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 
Chronic exposure to ethanol ± inhibitors and/or inducers of ethanol metabolism
Compared to acute ethanol exposures, long-term exposures significantly increased ethanol cytotoxicity. The percentage of viability was decreased in a time-dependent manner (P < 0.0001; Anova test). The longest periods of exposure were associated with the lowest rates of surviving cells (only 30% survival after 9 days) (Fig. 3A). Indeed, the Fisher PLSD test showed values different from controls, especially for the longest times of exposure (P < 0.0001). Within this time frame, DNA alterations were also noticeable after 3 and 6 days of ethanol treatment, reaching a steady level after 6 days (P < 0.0001; Fisher PLSD test; Fig. 3B).



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Fig. 3. (A) Astrocyte variation in viability after chronic exposures to 20 mM ethanol. Results are the mean of three independent experimental series and are expressed as mean ± SEM (n = 15) (***P < 0.0001 when groups exposed to ethanol are compared to control, Fisher PLSD test). Control without ethanol represents 100% viability. Experimental conditions are detailed in the Methods. (B) Level of DNA damage in rat astrocytes after chronic exposures to ethanol 20 mM and analysed by the comet assay. DNA damage is expressed as percentage of DNA in the tail. The figure shows mean results of three independent experiments. Fifty cells were randomly examined in triplicate for each condition and results are expressed as mean ± SEM (***P < 0.0001 when groups exposed to ethanol are compared to control, Fisher PLSD test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 
As 6 or 9 days of exposure induced very substantial cytotoxicity, a chronic exposure of 3 days was preferred for the experiments with inducers and/or inhibitors. Furthermore, methylene blue (MB) and glucose oxidase (GO) concentrations were decreased by 30 and 2, respectively, when compared to AI values, in order to minimize cytotoxicity due to these chemicals as much as possible.

Despite these lower concentrations, adding inhibitors and/or inducers to the culture medium without ethanol led to a general decrease in cell viability apart from 3-amino-1,2, 4-triazole (AMT) (GO vs control, P < 0.001; MB/GO vs control, P < 0.001; MB vs control, P < 0.0001 Mann–Whitney test) (Fig. 4A). Nevertheless, in presence of ethanol, the addition of these various inhibitors and/or inducers did not modify cell viability when compared to control with ethanol (control with ethanol vs various pre-treatments plus ethanol, NS Mann–Whitney test) (Fig. 4A).



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Fig. 4. (A) Astrocyte variation in viability after exposure to various pre-treatment in presence or absence of ethanol. Results are the mean of three independent experimental series and are expressed as mean ± SEM (n = 15). No statistical difference was observed when control exposed only to ethanol was compared to groups exposed to various pre-treatment plus ethanol, using the Fisher PLSD test. Control without ethanol represents 100% viability. Experimental conditions are detailed in the Methods. (B) Level of DNA damage in rat astrocytes exposed to various pre-treatment in presence or absence of ethanol. DNA damage is expressed as percentage of DNA in the tail. The figure shows mean results of three independent experiments. Fifty cells were randomly examined in quadruplicate for each condition and results are expressed as mean ± SEM (***P < 0.0001 when groups exposed to various pre-treatment plus ethanol were compared to group only exposed to pre-treatment, Mann–Whitney test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 
In the study of DNA damage, adding inhibitors and/or inducers in the culture medium without ethanol led to a significant rise in DNA strand breaks (MB vs control, P < 0.0001; GO vs control, P < 0.0001; MB/GO vs control, P < 0.0001; AMT vs control, P < 0.001, Mann–Whitney test) (Fig. 4B).

In presence of ethanol, the percentage of DNA in the tail was increasingly high, reflecting greater DNA damage under such conditions. The most severe impact was observed in circumstances corresponding to the most acetaldehyde-promoting conditions, i.e. after inhibition of aldehyde dehydrogenase by MB and catalase stimulation by GO. Indeed after adding ethanol the percentage of tail DNA was increased by more than 1.7 (MB/GO vs MB/GO + ethanol, P < 0.0001 Mann–Whitney test) (Fig. 4B). The distribution of the DNA tail values was also significantly modified in the presence of MB and GO and ethanol (MB/GO vs MB/GO + ethanol, P < 0.0001 Kolgomorov–Smirnov test) (Fig. 5). It should be noted that in presence of ethanol, the addition of MB/GO led to more DNA damage when compared to cells only exposed to ethanol (ethanol vs MB/GO + ethanol, P < 0.0001 Kolgomorov–Smirnov test) (Fig. 5). In this figure, an interesting change in the distribution of tail DNA can be observed. Indeed, the most abundant fraction of cells exposed to ethanol 20 mM had tail DNA between 0% and 10% (Fig. 5B), whereas the most abundant fraction of cells exposed to MB/GO plus ethanol had tail DNA between 40% and 70%, confirming the many DNA strand breaks.



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Fig. 5. Histograms of the distribution of comet tail DNA of astrocytes incubated with 20 mM ethanol for 3 days (CI) with or without MB/GO pre-treatment. (A) Control; (B) CI (B vs A, P < 0.0001, Kolgomorov–Smirnov test); (C) MB/GO (C vs A, P < 0.0001 and C vs B, P < 0.02, Kolgomorov–Smirnov test); (D) MB/GO plus CI (D vs A, D vs B, and D vs C, P < 0.0001, Kolgomorov–Smirnov test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 
Using AMT, a catalase inhibitor, a slight rise in DNA strand breaks was observed, whereas AMT plus ethanol (condition inducing ethanol accumulation) led to substantial DNA damage (Fig. 4B). Indeed, the percentage of tail DNA increased more than 1.6 times with AMT and ethanol compared to AMT alone (AMT + ethanol vs AMT, P < 0.0001 Mann–Whitney test) (Fig. 4B). The distribution of astrocytes exposed to AMT as a function of the percentage of tail DNA is shown in Fig. 6. In control cells (Fig 6A) and cells exposed to ethanol (Fig. 6B) or to AMT alone (Fig. 6C), the most abundant fraction of the cells had tail DNA between 0% and 20%. In cells exposed both to AMT and ethanol, we observed a significant increase in the percentage of cells with high tail DNA (between 30% and 50%) (Fig. 6D).



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Fig. 6. Histograms of the distribution of comet tail DNA of astrocytes incubated with 20 mM ethanol for 3 days (CI) with or without AMT pre-treatment. (A) Control; (B) CI (B vs A, P < 0.0001, Kolgomorov–Smirnov test); (C) AMT (C vs A, P < 0.001, C vs B, NS Kolgomorov–Smirnov test); (D) AMT plus CI (D vs A, D vs B, and D vs C, P < 0.0001, Kolgomorov–Smirnov test). Experimental conditions are detailed in the Methods. AU: arbitrary units.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The toxic action of ethanol has been studied intensively over the past years, but the underlying mechanisms of how the drug exerts its deleterious effects on development remains unknown (Brooks, 1997Go; Seitz et al., 1998Go). This is a major focus of investigation, especially with regard to the central nervous system, which is highly vulnerable to alcohol-induced damage. Several mechanisms have been proposed involving the enzyme-catalyzed oxidation of ethanol to acetaldehyde. Acetaldehyde is a highly reactive metabolite that may induce cytotoxic conditions by binding tissue macromolecules and disrupting cellular functions (Lieber, 1997Go). Furthermore, it was shown that free radicals were implicated in acetaldehyde metabolism (Gonthier et al., 1991Go; Nakao et al., 2000Go).

The level of DNA damage in a sample is the balance between the generation of DNA lesions and the repair of this damage if a repair pathway exists. It was reported that acetaldehyde decreases DNA repair mechanisms (Garro et al., 1992Go).

We chose to observe DNA damage using the comet assay because this method provides fast results and requires only a few cells. This technique therefore seems suitable for analysis in primary culture, which requires continual new preparations.

Whereas acute exposure to ethanol at all of the tested concentrations did not lead to loss of cell viability (data not shown), a chronic exposure induced a dose-dependent decrease in this viability (Fig. 3A). Similarly, acute ethanol exposure did not induce any DNA damage, as observed using the comet assay (data not shown), while chronic conditions led to a time-dependent rise in the percentage of DNA in the tail (Fig. 3B).

Acetaldehyde, the main metabolite of ethanol, can cause different cell modifications (Reinke et al., 1988Go; Wan et al., 2000Go; Niemela, 2001Go). It was therefore interesting to compare how acetaldehyde-induced DNA damage evolved with the damage generated by ethanol as a function of the doses of acetaldehyde or ethanol. Acetaldehyde doses were chosen according to the formation of acetaldehyde from ethanol metabolism as described by Holownia et al. (1996)Go. In their experiments, a constant acetaldehyde formation (450 µM) was obtained from 20 mM ethanol by alcohol dehydrogenase-transfected Chinese hamster ovary cells co-cultured with rat astrocytes. Other authors studied the effects of lower acetaldehyde concentration (50 µM), but they used sub-cellular fractions (i.e. synaptic membranes), an adequate model for evaluating the damaging effects on neurons (Chumakova et al., 2000Go).

As for acute ethanol exposure, no change in cytotoxicity was observed when cells were incubated with any of the acetaldehyde concentrations for 3 h (Fig. 1A). In contrast, single and double DNA strand breaks were enhanced in cells exposed to acetaldehyde in a dose-dependent manner (P < 0.0001, Anova test) (Fig. 1B).

Different authors also observed DNA damage after acetaldehyde exposure, either in co-cultured cells (Holownia et al., 1999Go) or in lymphocytes and gastrointestinal cells (Singh and Khan, 1995Go; Blasiak et al., 2000Go). However, their experimental conditions were more deleterious than ours, i.e. acetaldehyde concentrations were higher (up to 100 mM) and exposure times were longer (4 days).

The toxicity of acetaldehyde is confirmed by experiments done in the presence of inhibitors or inducers of ethanol metabolism. For example, the addition of glucose oxidase (GO), a known hydrogen peroxide generator, to the incubation medium significantly increases the generation of acetaldehyde in homogenates of perfused rat brains incubated in the presence of ethanol (50–100 mM) (Aragon et al., 1992Go; Eysseric et al., 2000Go). Moreover, methylene blue (MB) can be used to inhibit aldehyde dehydrogenase (ALDH), so it accumulates acetaldehyde in the cell (Helander et al., 1993Go; Eysseric et al., 2000Go).

In acute ethanol conditions, the presence of MB and/or GO resulted in no modification in either the percentage of cytotoxicity or in the percentage of DNA in the tail (data not shown). In chronic exposure, the pre-treatment with MB and/or GO, in absence of ethanol, induced a substantial increase in cell toxicity, despite lower concentrations compared to acute exposure. It was interesting to note that in presence of alcohol, none of the pre-treatments, whatever their nature, resulted in a significant modification of viability compared to controls with ethanol (Fig. 4A).

When the acetaldehyde dehydrogenase was inhibited by MB and the catalase was stimulated by GO, the addition of ethanol led to a very high increase in the percentage of tail DNA, indicating substantial DNA damage (Fig. 4B). This observation could be explained by an accumulation of acetaldehyde in cells under these conditions (Eysseric et al., 2000Go). This pre-treatment with MB/GO provides evidence for the toxic role of acetaldehyde, the main metabolite of ethanol oxidation, and confirms results obtained by Holownia et al. (1999)Go. These results were further supported by significant modifications in the distribution of the percentage of DNA in the tail between cells exposed only to ethanol (Fig. 5B) or to MB/GO (Fig. 5C) and cells exposed to ethanol with MB/GO (Fig. 5D).

We also tested 3-amino-1,2,4-triazole (AMT), an inhibitor of catalase. This is the main enzyme in the brain catalysing the transformation of ethanol to acetaldehyde (Aragon et al., 1992Go; Gill et al., 1992Go). Indeed, inhibition of catalase levels has been known to reduce the rate of cerebral oxidation of ethanol and thus favour ethanol accumulation within the cell (Aragon and Amit, 1992Go; Hamby-Mason et al., 1997Go). Adding AMT in the culture medium led to no significant increase in cell cytotoxicity or DNA damage formation during acute exposure (data not shown). Nevertheless, it is interesting to note that in chronic exposure, the percentage of DNA in the tail was higher with ethanol and AMT than with ethanol alone (P < 0.0001, Mann–Whitney test) (Fig. 4B). These results were confirmed by significant modifications in the distribution of DNA in the tail (Fig. 6). Indeed, after exposure to AMT and ethanol, the percentage of damaged cells greatly increased and the percentage of tail DNA shifted towards higher values when compared to controls without ethanol or to chronic exposure.

In brain, catalase has a dual action: on one hand it generates acetaldehyde from ethanol; on the other hand it plays a protective role against oxidative stress (Harris, 1992Go; Islam et al., 1997Go). The use of AMT, which inhibits this enzyme, leads not only to an alcohol accumulation in the astrocytes, but also to an increase in intracellular H2O2, the single cofactor of catalase. Recent investigations have found that an important ethanol concentration in astrocytes cannot be entirely metabolized by the cells, so the excess of ethanol could function as a hydroxyl radical scavenger and play a role in a detoxification process (Russo et al., 2001Go). Moreover, our conditions of chronic ethanol exposure promote the induction of cytochrome P450 2E1 (Anandatheerthavarada et al., 1993Go; Brzezinski et al., 1999Go; Dupont et al., 2000Go; Upadhya et al., 2000Go). This pathway is known to induce in brain both the formation of reactive oxygen species (Rashba-Step et al., 1993Go; Bondy and Oroszo, 1994Go) and the formation of {alpha}-hydroxyethyl radicals derived from ethanol (Gonthier et al., 1991Go; Gonthier et al., 1997Go). In addition, these free radicals produced during ethanol metabolism induce DNA damage (Gonthier et al., in press) and inhibit various antioxidant enzymes (e.g. catalase, SOD, GPX), thus increasing cell damage (Puntarulo et al., 1999Go). Moreover, the enhancement of the hydrogen peroxide concentration through catalase inhibition promotes this free radical pathway, adding to cellular damage such as DNA strand breaks.

Finally, the large increase in DNA damage observed after AMT addition seems to have been caused by a deleterious effect of free radicals and a concomitant decrease in antioxidant protection rather than by a toxic action of the non-metabolized ethanol in the cells. Nevertheless, we cannot exclude the potential damaging effect on DNA of this non-metabolized ethanol, especially in these conditions favouring free radical formation but also ethanol accumulation; however, precisely evaluating how extensive this may be appears problematic.

These results confirm the hypothesis that processes other than catalase activity contribute to brain ethanol metabolism (Zimatkin et al., 1998Go). Moreover, in previous studies conducted on brain microsomes, we provided evidence that ethanol metabolism via the MEOS pathway (an enzymatic system containing cytochrome P-4502E1) led to {alpha}-hydroxyethyl radical formation (Gonthier et al., 1991Go). Other research excludes CYP2E1 as a possible second pathway because the inhibition of this enzyme had no effect on acetaldehyde production (Gill et al., 1992Go). Nevertheless, it should be noted that these investigators used metyrapone, a non-specific inhibitor of cytochromes P-450. In order to study this question thoroughly, we suggest investigating this possibility using a specific inhibitor of CYP2E1 such as diallyl sulphide.

In summary, at low doses, acute ethanol exposure of astrocytes produces no effect on either cell viability or DNA strand breaks. However, acute exposure to acetaldehyde leads to DNA strand breaks despite no apparent effect on cell viability. On the other hand, chronic exposure to ethanol induces both DNA alterations and loss of cell viability, which are raised by the concomitant use of MB and GO leading to an intracellular accumulation of acetaldehyde. In conditions favouring both the metabolism of ethanol by a free radical pathway and ethanol accumulation (i.e. inhibition of catalase by aminotriazole), DNA damage is also observed but to a lesser extent than in conditions promoting acetaldehyde formation.

These experimental findings could thus favour the hypothesis that acetaldehyde plays a leading role in the generation of the toxic effects on cells after ethanol exposure. Nevertheless, a direct effect of ethanol on DNA damage cannot be ruled out, especially when the cell is chronically exposed to ethanol.


    ACKNOWLEDGEMENTS
 
This work was supported by the Institut de Recherches Scientifiques sur les Boissons (IREB).


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Anandatheerthavarada, H. K., Shankar, S. K., Bhamre, S., Boyd, M. R., Song, B. J. and Ravindranath, V. (1993) Induction of brain cytochrome P-450IIE1 by chronic ethanol treatment. Brain Research 601, 279–285.[CrossRef][ISI][Medline]

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Aragon, C. M., Rogan, F. and Amit, Z. (1992) Ethanol metabolism in rat brain homogenates by a catalase–H2O2 system. Biochemical Pharmacology 44, 93–98.[CrossRef][ISI][Medline]

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