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)
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ABSTRACT |
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INTRODUCTION |
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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, 1989). Astrocytes may also be a major site for the detoxification or the bioactivation of neurotoxins (Di Monte et al., 1996
).
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) used ethanol concentrations of 50, 100 and 200 mM for 10 days, whereas Renis et al. (1996)
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.
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METHODS |
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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, 1972), with a few modifications (Gonthier et al., 1997
).
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 -GFAP, an astrocyte marker (Vijayan et al., 1993
).
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., 1997).
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., 1992). 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., 1996
). 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., 1993
).
Determination of cell viability
The viability of the astrocytes after different stresses was determined using the MTT reduction test (Iselt et al., 1989). 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., 1993) 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). 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 TrisHCl, 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, 516560 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, 2003).
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 MannWhitney test when appropriate. The KolgomorovSmirnov test was employed to compare the distribution of the percentages of DNA in the tail.
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RESULTS |
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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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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 MannWhitney 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 MannWhitney test) (Fig. 4A).
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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 MannWhitney 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 KolgomorovSmirnov 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 KolgomorovSmirnov 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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DISCUSSION |
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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., 1992).
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., 1988; Wan et al., 2000
; Niemela, 2001
). 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)
. 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., 2000
).
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., 1999) or in lymphocytes and gastrointestinal cells (Singh and Khan, 1995
; Blasiak et al., 2000
). 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 (50100 mM) (Aragon et al., 1992; Eysseric et al., 2000
). Moreover, methylene blue (MB) can be used to inhibit aldehyde dehydrogenase (ALDH), so it accumulates acetaldehyde in the cell (Helander et al., 1993
; Eysseric et al., 2000
).
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., 2000). 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)
. 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., 1992; Gill et al., 1992
). 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, 1992
; Hamby-Mason et al., 1997
). 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, MannWhitney 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, 1992; Islam et al., 1997
). 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., 2001
). Moreover, our conditions of chronic ethanol exposure promote the induction of cytochrome P450 2E1 (Anandatheerthavarada et al., 1993
; Brzezinski et al., 1999
; Dupont et al., 2000
; Upadhya et al., 2000
). This pathway is known to induce in brain both the formation of reactive oxygen species (Rashba-Step et al., 1993
; Bondy and Oroszo, 1994
) and the formation of
-hydroxyethyl radicals derived from ethanol (Gonthier et al., 1991
; Gonthier et al., 1997
). 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., 1999
). 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., 1998). 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
-hydroxyethyl radical formation (Gonthier et al., 1991
). Other research excludes CYP2E1 as a possible second pathway because the inhibition of this enzyme had no effect on acetaldehyde production (Gill et al., 1992
). 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.
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ACKNOWLEDGEMENTS |
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