IN VITRO EFFECTS OF ETHANOL, ACETALDEHYDE AND FATTY ACID ETHYL ESTERS ON HUMAN ERYTHROCYTES

O. V. Tyulina, V. D. Prokopieva1, R. D. Dodd2, J. R. Hawkins2, S. W. Clay3, D. O. Wilson2, A. A. Boldyrev and P. Johnson2,4,*

International Biotechnological Center and Center for Molecular Medicine of MV Lomonosov, Moscow State University, 119899 Moscow,
1 Mental Health Research Institute, Medical Academy of Sciences of Russia, Tomsk, Russia,
2 Department of Chemistry and Biochemistry,
3 Department of Geriatric Medicine and
4 Department of Biomedical Sciences, Ohio University, Athens, OH 45701, USA

Received 24 April 2001; in revised form 27 July 2001; accepted 23 August 2001


    ABSTRACT
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
In vitro experiments were performed to determine if ethanol was metabolized by human erythrocytes and to investigate if ethanol or its metabolites, acetaldehyde and fatty acid ethyl esters, affected erythrocyte morphology and stability. No detectable metabolism of ethanol was found in erythrocytes, although ethanol itself caused an elevated rate of spontaneous haemolysis in erythrocyte preparations. Physiologically attainable levels of ethanol were found to stabilize erythrocytes against haemolysis induced by sodium hypochlorite, and the presence of ethanol caused a decrease in erythrocyte reactive oxygen species levels, although the mechanism for such a process is unknown. Both physiologically attainable and higher levels of acetaldehyde had no effects on erythrocyte morphology and stability even after a 16 h exposure. Fatty acid ethyl esters caused structural changes and instability in erythrocytes in vitro, but whether such changes occur in vivo has not been established. The results of these studies suggest that the deleterious effects of ethanol consumption on erythrocytes in vivo may be, at least in part, the result of direct effects of unmetabolized ethanol on erythrocyte components.


    INTRODUCTION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
In a variety of animal tissues and cells, the principal route of ethanol metabolism is its oxidation to acetaldehyde by alcohol dehydrogenase, ethanol-inducible cytochrome P-450 (CYP2E1), and catalase (Lieber, 1994Go; Chen et al., 1995Go; Kishimoto et al., 1995Go; French, 1996Go). Such metabolism is now recognized as being potentially harmful at high levels, because these oxidative processes can generate the hydroxyethyl radical, which can cause cell damage and cell death in some cases (Reinke et al., 1997Go). Additionally, the oxidation of ethanol to acetaldehyde is deleterious because acetaldehyde is a highly reactive species which can modify proteins and other molecules (Hipkiss et al., 1998Go; Holownia et al., 1999Go). In other tissues and cell types, such as nerve cells and cardiomyocytes, non-oxidative conversion of ethanol to fatty acid ethyl esters by fatty acid ethyl ester synthases appears to be the predominant pathway of ethanol metabolism (Gorski et al., 1999Go; Laposata, 1999Go). These esters appear to cause cell damage by interfering with several different aspects of cell metabolism including membrane transport functions (Gubitosi-Klug and Gross, 1996Go). In addition to the effects of ethanol metabolites, unmetabolized ethanol may also have direct effects on cells, and it has been shown that it can permeate membranes and cause disruption of normal cell structure and metabolism (Trandum et al., 1999Go; Wirkner et al., 1999Go).

In the case of erythrocytes, these cells have been shown to lack significant activities of fatty acid ethyl ester synthases, alcohol dehydrogenase or CYP2E1, although they do contain high catalase activity (Zorzano et al., 1989Go). Exposure in vivo and in vitro to ethanol has been shown to affect a number of erythrocyte properties, including cell morphology, in vivo lifetime and resistance to haemolysis (Prokopieva et al., 2000Go). The cause of this damage has not been clearly delineated, but could come from direct effects of ethanol on the cell, the production of acetaldehyde inside the erythrocyte by catalase (Tyulina et al., 2000Go) or the incorporation into the cells of fatty acid ethyl esters synthesized in other cell types, but transiently present in the circulatory system following alcohol consumption (Doyle et al., 1994Go).

In order to assess which of these possible mechanisms is responsible for ethanol-related damage in human erythrocytes, we have conducted in vitro studies in which we have measured free radical levels in these cells as a result of their exposure to ethanol and acetaldehyde, the effects of inhibition of erythrocyte catalase, and the effects of the presence of fatty acid ethyl esters.


    MATERIALS AND METHODS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Materials
Dichlorodihydrofluorescein diacetate (DCDFDA) and fluorescein isothiocyanate (FITC)-labelled annexin V were purchased from Molecular Probes (Eugene, OR, USA). The Vac-Elut apparatus and solid-phase extraction columns (Bond Elut NH2; 500 mg, 3 ml capacity) were products of Varian (Harbor City, CA, USA). The BCA protein concentration reagents were purchased from Pierce (Rockford, IL, USA), and 4–6% sodium hypochlorite (NaOCl) solution was a product of Fisher Chemicals (Fair Lawn, NJ, USA). Hanks' balanced salt solution (item H4891), fatty acid ethyl esters, 3-aminotriazole, and methylene blue were obtained from Sigma Chemical Co. (St Louis, MO, USA).

Erythrocyte sources and preparation
For in vitro experiments on isolated erythrocytes at Ohio University, cells were donated by three alcohol-abstinent adult males in normal health. At least three different erythrocyte preparations were used in all the experiments. Institutional approval for the use of human subjects in this study was obtained from Ohio University, and informed consent of all donors was obtained prior to blood sample collection. Sodium citrate (final concentration 0.38% w/v) was used to prevent coagulation of the drawn blood and erythrocyte purification was performed as previously described (Chi and Wu, 1991Go) using Hanks' balanced salt solution (HBSS: 4.2 mM NaHCO3, 5 mM KCl, 0.4 mM KH2PO4, 138 mM NaCl, 0.34 mM Na2HPO4, 5 mM glucose, pH 7.0) as the wash buffer. Erythrocyte preparations were made within 1 h of blood collection and experiments were performed within 4 h of erythrocyte preparation.

Haemolysis assays
For measurement of oxidatively-induced haemolysis, either 0.1 mM cumene hydroperoxide (Yesilkaya et al., 1998Go) from a fresh cold 100 mM stock solution in dimethylsulphoxide (DMSO) or 0.03 mM NaOCl (Cherny et al., 1992Go) from a fresh cold 7.5 mM (determined at pH 12.0 from the molar absorbance at 290 nm of 350 M–1 cm–1) stock solution in water were used in 2 ml samples of erythrocytes which contained ~107 cells (corresponding to 0.7 units of absorbance at 630 nm). Haemolysis at 23 ± 0.5°C in HBSS solution, pH 7.0 was initiated by addition of the oxidizer and the progress of haemolysis was monitored at 630 nm. Haemolysis was measured until no further changes in absorbance were registered; for data analysis, the difference between initial and final absorbance values was designated as 100% haemolysis. Verification that total haemolysis had occurred was seen as there were no further absorbance changes on addition of HCl at a 1 M final concentration, a condition known to cause total haemolysis (Ivanov, 1999Go). The value of T50 (the time at which 50% of the cells were haemolysed) was calculated as decreased values of T50 reflect increased susceptibility to haemolysis (Prokopieva et al., 2000Go). Duplicate or triplicate analyses of haemolysis curves for each sample were performed.

For experiments on the effects of ethanol and acetaldehyde on spontaneous erythrocyte haemolysis in the absence of added oxidizing agent, erythrocyte preparations at 40% haematocrit were shaken gently at 25°C for 16 h in the presence of different concentrations of ethanol, up to 0.5% (v/v). Haemolysis levels in the presence of ethanol were compared to levels in cells incubated in the absence of ethanol. For similar experiments on the effects of fatty acid ethyl esters (FAEE), erythrocyte preparations at 40% haematocrit in the absence or presence of 40 mg/ml of human serum albumin were gently shaken at 25°C for 24 h in the absence or presence of 100 µM linolenyl (LEE) or palmityl (PEE) ethyl ester, added from stock 10 mM solutions in DMSO. Haemolysis levels in the presence of FAEE were compared to levels in control samples that contained no FAEE but contained the same amount of DMSO. Gentamycin (0.01 mg/ml), penicillin, streptomycin (5 units/ml each) and fungizone (0.025 µg/ml) were added to the samples as antibiotics (Huentelman et al., 1999Go). In these experiments, the total extent of haemolysis that had occurred during the incubation period was calculated from the ratio of haemoglobin (determined by absorbance measurement at 415 nm) in the supernatants of the samples after centrifugation of cells compared to that in a haemolysate of the same erythrocyte sample prepared by dilution of the sample into 10 vol of 5 mM phosphate buffer, pH 8.0.

Measurement of reactive oxygen species (ROS) levels in erythrocytes
Measurement of ROS levels inside erythrocytes was performed by flow cytometric analysis of intracellularly generated 2',7'-dichlorofluorescein (DCF) as described previously (Johnson et al., 1998Go). Erythrocyte samples were diluted in HBSS to ~106 cells/ml and the cells were pre-incubated at 37°C for 30 min in the dark. 2',7'-Dichlorodihydrofluorescein diacetate (DCDFDA) was then added in the dark to a final concentration of 100 µM using a stock 10 mM solution in DMSO, and the cells were incubated for 1 h, after which ethanol additions were made to cell samples to final concentrations of 0.1 and 0.5%. These and no-ethanol control samples were then incubated at 25°C for further periods of time up to 16 h, after which 104 cells from the samples were analysed on a Becton–Dickinson FACSort Flow Cytometer equipped with Lysis II software.

Inhibition of erythrocyte catalase activity and enzyme activity assays
Inhibition of catalase activity was carried out by incubation of washed erythrocytes at ~108 cells/ml in HBSS, which contained 3-aminotriazole (25 mM) and methylene blue (0.01 mg/ml) for 60 min at 37°C (Tephly et al., 1961Go). In order to remove 3-aminotriazole and methylene blue, cell samples were centrifuged after treatment at 3000 g for 5 min, resuspended in 6 volumes of HBSS and incubated for 15 min at 37°C. This wash procedure was repeated four times, and the erythrocytes were then suspended in HBSS at ~108 cells/ml before further use.

Analysis of catalase activity in cell samples was performed by the assay procedure of Aebi (1984). In comparison to control cells untreated with 3-aminotriazole or methylene blue, treated cells showed an inhibition of catalase activity of 95–100%. Assays of the erythrocyte antioxidant enzymes superoxide dismutase and glutathione peroxidase were also performed following the catalase inactivation procedure to verify that these enzyme activities were unaffected by this treatment. In these assays, superoxide dismutase activity was measured by the cytochrome c reduction procedure in the absence of cyanide, in order to measure total superoxide dismutase activity (Paoletti and Mocali, 1990Go), and glutathione peroxidase was assayed by the glutathione reductase-linked ‘continuous assay’ procedure of Flohe and Gunzler (1984).

Determination of ethanol utilization by erythrocytes
In experiments to determine if erythrocytes metabolized ethanol, the concentration of ethanol in the medium before and after incubation in the presence of 109 erythrocytes/ml of HBSS containing between 0.05 and 0.5% ethanol was determined by headspace gas chromatography (Meyer, 1978Go). Before analysis, samples were centrifuged to remove cells and 2 ml aliquots were transferred into 10 ml headspace vials. For analysis, triplicate samples of headspace vapour were injected onto a Restek RPX-624 column (30 m x 0.53 mm ID, 3 µm film) on an HP 4890 gas chromatograph interfaced with HP ChemStation software. Under the conditions of operation (He carrier gas, 35°C isothermal column temperature), acetaldehyde and ethanol eluted at ~0.7 min and 1.4 min respectively, and isopropanol, which was added to sample aliquots as an internal recovery standard, eluted at 1.8 min.

Analysis of FAEE contents of erythrocyte membranes
Erythrocyte membrane fractions from erythrocytes exposed and unexposed to FAEE were prepared as described by Akoev et al. (1998) and protein contents of the preparations were determined by the BCA reagent. Triplicate aliquots of membrane samples based on equivalent protein contents were then prepared for gas chromatographic analysis by a solid phase extraction procedure (Bird et al., 1997Go). For gas chromatographic analysis of the extracts, a Supelco SPB-1 column (15 m x 0.25 mm ID, 1 µm film) installed on a Hewlett Packard 5890 gas chromatography system equipped with HP ChemStation software was used. Under the conditions of operation (He carrier gas, 180°C isothermal column temperature), the approximate elution times of the ethyl esters of palmitate, oleate, linolenate and stearate were 22, 42, 43 and 49 min respectively.

Phosphatidylserine (PS) distribution analysis
Isolated erythrocytes in HBSS were incubated for 24 h at 25°C in the presence of 100 µM LEE or PEE. The erythrocytes were then transferred to HBSS containing 1.5 mM Ca2+ and were treated with 0.06 µg/ml FITC-labelled annexin V at a cell density of 106/ml for 30 min at 25°C according to the procedures described by Kuypers et al. (1996). After washing in HBSS containing 2.5 mM Ca2+, the cells were immediately analysed to measure the extent of PS externalization as determined by FITC–annexin V binding to the cells (Koopman et al., 1994Go) using 104 cells from triplicate samples for analysis on a Becton–Dickinson FACSort Flow Cytometer equipped with Lysis II software. In these studies, cells treated with DMSO but not exposed to FAEE were used as the negative control cells, and a positive control for the maximum externalization of PS in the cells was prepared by treatment of cells with Ca2+, a calcium ionophore and N-ethylmaleimide.

Statistical analysis
The data in the tables and figures are presented as means ±SD (bars), and one-way ANOVA using the Dunnett method was performed to determine significant differences between data sets. Data sets with P < 0.05 were considered to be significantly different.


    RESULTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
Effects of exposure of human erythrocytes to ethanol
When erythrocytes were incubated in the presence of ethanol for 16 h, it was found that there was a small, but significant dose-dependent increase in haemolysis at ethanol concentrations of >=0.3%, as determined by the release of haemoglobin into the medium (Fig. 1Go). The extent of haemolysis in the absence of ethanol was found to be 0.66%, whereas that in the presence of 0.5% ethanol was 1.11%. Microscopic examination of the erythrocytes incubated in the absence or presence of ethanol revealed no obvious differences in morphology (results not shown).



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Fig. 1. Haemolysis of erythrocytes during incubation with ethanol.

The data mean ± SD (bars) show results of experiments in which erythrocytes at a 40% haematocrit were incubated with different concentrations of ethanol for 16 h at 25°C. Data were obtained from six separate experiments with triplicate assays of haemolysis performed as described in the Materials and methods section. The asterisk indicates that the value is significantly different from the control (no ethanol) value in which the extent of haemolysis was 0.72 ± 0.04%.

 
In further experiments, erythrocytes were incubated in the absence and presence of ethanol and then analysed for their resistance to haemolysis induced by NaOCl (Fig. 2Go). It was found that the resistance of the cells to oxidatively-induced haemolysis was significantly increased compared to the control value when incubation was performed in the presence of >=0.2% ethanol concentrations, with erythrocytes incubated in 0.5% ethanol having an average T50 time 2.17-fold that of the control (no ethanol) cells.



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Fig. 2. Effects of ethanol on T50 values of erythrocytes.

The data mean ± SD (bars) show results of experiments in which erythrocytes at a 40% haematocrit were incubated with different concentrations of ethanol for 16 h at 25°C and then subjected to haemolysis by 0.03 mM NaOCl. Data were obtained from six separate experiments with triplicate assays of haemolysis performed as described in the Materials and methods section. The asterisk indicates that the value is significantly different from the control (no ethanol) value for which the T50 value was 2.55 ± 0.26 min.

 
Because the production of free radicals is known to be involved in the mechanism of NaOCl-induced haemolysis (Panasenko and Arnhold, 1996Go), the results from our NaOCl-induced haemolysis studies, which showed that ethanol can stabilize cells against NaOCl effects, suggested that the presence of ethanol may decrease free radical levels in the erythrocytes. In order to investigate this further, analysis of ROS levels inside the cells in the absence and presence of 0.5% ethanol was performed by the DCF procedure. A representative flow cytometric profile is shown in Fig. 3Go, with experimental data shown in Fig. 4Go. These results indicate that ROS levels were significantly decreased in the presence of ethanol, in comparison with levels observed in cells incubated in the absence of ethanol.



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Fig. 3. Effects of ethanol on the 2',7'-dichlorofluorescein fluorescence of erythrocytes.

Representative curves are shown for erythrocytes pre-incubated at 106 cells/ml in the absence and presence of 0.5% (v/v) ethanol for 16 h at 25°C. 2',7'-Dichlorofluorescein (DCF) fluorescence intensity in arbitrary units (horizontal axis) is plotted on a logarithmic scale against cell numbers on a linear scale.

 


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Fig. 4. Effects of ethanol (EtOH) on the 2',7'-dichlorofluorescein (DCF) fluorescence of erythrocytes with active and inactivated catalase.

The bar charts show data mean ± SD (bars) for experiments in which DCF fluorescence was measured in erythrocyte cell preparations pre-incubated in the absence or presence of 0.5% (v/v) ethanol for 16 h at 25°C following treatment (or no treatment) to inactivate erythrocyte catalase activity to <5% of control value. The data are expressed relative to the DCF fluorescence level in the control (no ethanol, active catalase) cells. The asterisk indicates that the value is significantly different from the control value.

 
Studies on erythrocytes with inactivated catalase activity
The significance of the results from the previous section is that they suggest that erythrocyte structural changes and haemolysis caused by ethanol in vivo may not be caused by oxidative metabolism of ethanol in the erythrocyte. In order to test this hypothesis, further studies were performed on erythrocytes in which catalase activity had been inactivated, as catalase is the only major erythrocyte enzyme with the potential for ethanol oxidation (Zorzano et al., 1989Go; Hamby-Mason et al., 1997Go).

On analysis by the DCF procedure (Fig. 4Go), it was found that erythrocytes with inactivated catalase activity had significantly increased ROS levels (141% of the control cell value), and analysis of the resistance of these cells to oxidative haemolysis induced by cumene hydroperoxide (Fig. 5Go), as measured by the T50 haemolysis time (90% of control value), revealed that they were significantly more susceptible to haemolysis than were cells with normal catalase activity.



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Fig. 5. Effects of ethanol on T50 values of human erythrocytes with active and inactivated catalase.

The data mean ± SD (bars) show results of experiments in which erythrocytes with active and inactivated catalase at a 40% haematocrit were incubated in the absence and presence of 0.5% (v/v) ethanol for 16 h at 25°C and then subjected to haemolysis by 0.1 mM cumene hydroperoxide. The asterisk indicates that the value is significantly different from the control (no ethanol, active catalase) value for which the T50 value was 59.0 ± 3.5 min.

 
When erythrocytes with inactivated catalase were incubated with ethanol, ROS levels were found to be significantly lower than those in no-ethanol catalase-inhibited control cells, but higher than those in cells with active catalase (Fig. 4Go). Although the presence of ethanol in catalase-inactivated cells decreased ROS levels in these cells, it caused no significant difference in the T50 haemolysis time for the catalase-inactivated cells (Fig. 5Go).

Determination of ethanol utilization by human erythrocytes
The conclusion that ethanol is not metabolized by catalase or other erythrocyte enzymes to a detectable extent was further substantiated by gas chromatographic analysis of the ethanol contents of the supernatants of erythrocyte preparations incubated in the absence and presence of between 0.05 and 0.5% ethanol. As shown in Table 1Go, these analyses revealed that no significant decreases in ethanol concentration occurred during the incubation period, thus indicating that there was no detectable metabolism of ethanol by the erythrocytes within the limits of the experimental technique.


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Table 1. Erythrocytes do not metabolize ethanol in Hanks' balanced salt solution
 
Effects of exposure of human erythrocytes to acetaldehyde
Although the above results indicate that substantial proportions of ethanol are not converted to acetaldehyde in erythrocytes, this does occur in a number of other tissues, (principally liver), and gives rise to a low (up to 2 µM) circulatory level of acetaldehyde consequent to ethanol ingestion (Green and Baron, 1986Go; Zorzano and Herrara, 1989). In order to determine if such exogenously produced acetaldehyde could affect erythrocyte morphology and stability, experiments were performed with erythrocytes in the presence of a physiologically attainable level of acetaldehyde (2 µM) that could be produced from the oxidative metabolism of ethanol in other tissues. Using the same conditions as for the experiments involving ethanol, this concentration of acetaldehyde was found to have no significant effect on erythrocytes in terms of the extent of spontaneous haemolysis (0.75 ± 0.07 and 0.66 ± 0.08% in the presence and absence of acetaldehyde), the resistance to oxidative haemolysis induced by NaOCl (measured T50 times were 4.22 ± 0.29 and 4.18 ± 0.40 min in the presence and absence of acetaldehyde, respectively), and cell morphology (results not shown). In additional experiments performed using a much higher concentration of acetaldehyde (3.6 mM), there was still no significant effect on the extent of spontaneous haemolysis (0.75 ± 0.07 and 0.82 ± 0.09% in the presence and absence of acetaldehyde), on the T50 values (4.30 ± 0.68 and 4.25 ± 0.62 min in the presence and absence of acetaldehyde), and on erythrocyte morphology (results not shown).

Effects of fatty acid ethyl esters on human erythrocytes
The above studies have shown that ethanol can have deleterious effects on erythrocyte stability as measured by the extent of haemolysis, although there is no detectable oxidative metabolism of ethanol by erythrocytes. Further studies were then performed on the effects of fatty acid ethyl esters on erythrocyte stability and structure as these compounds can be synthesized by a variety of cells in humans and are transiently present in the circulatory system (Doyle et al., 1994Go).

When erythrocytes at a physiological haematocrit level (40%) were incubated with physiologically attainable levels (100 µM) of two of the common fatty acid ethyl esters (LEE and PEE), it was found that incorporation of both esters into the erythrocyte membrane fraction occurred (Table 2Go).


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Table 2. Fatty acid ethyl ester (FAEE) levels in erythrocyte membranes following in vitro exposure to 100 µM FAEE
 
Analysis of the extent of spontaneous haemolysis in these preparations showed that both esters caused significant increases in the extent of haemolysis following incubation with the esters (Fig. 6Go), indicating that they both had disruptive effects on the cell membrane. When these experiments were performed in the presence of physiological levels (40 mg/ml) of albumin, a protein which binds FAEE in the blood (Bird et al., 1997Go), the destabilizing effects of the FAEE on haemolysis were greatly reduced but remained significant in comparison to control cells (Fig. 6Go).



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Fig. 6. Effects of fatty acid ethyl esters on erythrocyte haemolysis.

The bar charts show the data mean ± SD (bars) for the effects of 100 µM linolenyl (LEE) and palmityl (PEE) ethyl ester on the extent of haemolysis of isolated erythrocytes after 24 h. The shaded bars show results for experiments performed in the absence of albumin in the medium and the unshaded bars show results obtained in the presence of 40 mg/ml of albumin. The data are expressed relative to the haemolysis level in the control (no ester) cells, which was 0.64 ± 0.04%. The asterisk indicates that the value is significantly different from the control (no ester) value.

 
As it is known that decreased erythrocyte stability can occur as a result of PS externalization in the erythrocyte membrane (Diaz et al., 1996Go; Kuypers et al., 1998Go), the possibility that the lipophilic FAEE may cause such a change was investigated by flow cytometric analysis. As shown in Fig. 7Go, these studies revealed that both LEE and PEE caused small but significant elevations (14 and 4% respectively) in PS externalization, and when the erythrocytes were incubated with the esters in the presence of albumin, significant externalization still occurred. The extent of membrane PS externalization observed in these experiments was, however, small in comparison to the maximum possible externalization achievable in the presence of N-ethylmaleimide and a calcium ionophore (Kuypers et al., 1996Go), which was ~100-fold greater than that seen with the FAEE. Examination (results not shown) by light microscopy of FAEE-exposed erythrocyte preparations which showed increased haemolytic susceptibility and PS externalization indicated that these cells showed no indications of the abnormal morphology observed in erythrocytes from alcoholics (Prokopieva et al., 2000Go).



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Fig. 7. Effects of fatty acid ethyl esters (FAEE) on erythrocyte membrane phosphatidylserine distribution.

The bar charts show the data (mean ± SD) for the effects of 100 µM linolenyl (LEE) or palmityl (PEE) ethyl ester on fluorescein isothiocyanate–annexin-V fluorescence intensity after a 24 h incubation in Hanks' balanced salt solution in the absence (shaded bars) and presence (unshaded bars) of 40 mg/ml albumin. The results are normalized to a fluorescence intensity of 100 arbitrary fluorescence units in the negative control (no ester) value. The asterisk indicates that the value for fluorescence intensity in the presence of ester was significantly higher than the control value.

 

    DISCUSSION
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
The results from the haemolysis studies on the effect of ethanol on erythrocyte stability indicate that ethanol has a destabilizing effect on erythrocytes, and confirm similar conclusions from previous studies (Chi and Wu, 1991Go). Visual examination of the ethanol-exposed erythrocytes revealed no obvious morphological changes in the preparations, although it is known that after longer periods of exposure to ethanol, morphological changes can occur (McLawhon et al., 1987Go). It is therefore possible that the lack of morphological change under our experimental conditions was a reflection of the relatively small proportion of cells damaged (<1% as determined by haemolysis) during the incubation period used.

In contrast to these results, our previous studies (Tyulina et al., 2000Go) on rabbit erythrocytes have shown that, in unmodified cells, NaOCl-induced haemolysis was not retarded in the presence of ethanol, although ethanol had a stabilizing effect on haemolysis in catalase-inactivated cells. These results led us to suggest that oxidative metabolism of ethanol to acetaldehyde occurred in rabbit erythrocytes, whereas the present studies indicate that undetectable amounts of ethanol are metabolized by this enzyme and suggest that catalase does not play a major role in ethanol oxidation in the human erythrocyte, at least in the time periods of the in vitro experiments. Although the catalase activities of erythrocytes from humans and rabbits are comparable when assayed with H2O2 (Kurata et al., 1993Go), it is possible that catalase in the human erythrocyte has a much lower affinity for ethanol than the rabbit enzyme. Our studies on human catalase do, however, support previous conclusions that catalase functions as an important antioxidant enzyme which is involved in H2O2 removal and maintenance of erythrocyte stability (Halliwell and Gutteridge, 1999Go), as we have shown that erythrocytes are destabilized when catalase is inhibited in the absence of ethanol.

Circulatory acetaldehyde generated in vivo from ethanol metabolism has a short (3 h) lifetime (Zorzano and Herrara, 1989), and the erythrocyte itself contains an enzyme, aldehyde dehydrogenase (Wierzchowski et al., 1997Go), which is capable of oxidizing acetaldehyde to acetate and presumably minimizing acetaldehyde-induced damage to the cell. Our experiments with physiologically attainable (and even higher) levels of acetaldehyde show that this compound does not significantly affect erythrocyte morphology and stability over the time period (16 h) of the experiments. These combined observations suggest that erythrocyte damage that occurs in vivo resulting from non-chronic ethanol consumption is probably not a consequence of exposure of the erythrocytes to exogenously produced acetaldehyde, or from small amounts of ethanol that could be metabolized to acetaldehyde in the erythrocyte itself. However, in chronic ethanol drinking, prevailing acetaldehyde levels would be maintained for longer periods and could conceivably cause damage which was not detectable in our in vitro experiments.

The seeming paradox between the direct haemolytic effect of ethanol on erythrocytes (Fig. 1Go) and the stabilizing effect of ethanol on erythrocytes undergoing NaOCl-induced haemolysis (Fig. 2Go) could be explained by the relatively small destabilizing effect of ethanol which is observed (<1% haemolysis) over 16 h. This effect would be negligible in the short time period (generally <10 min) assay for NaOCl-induced haemolysis where 100% of the cells are haemolysed. An alternative explanation for this paradox is that the mechanisms of haemolysis induced by ethanol and NaOCl are different. The fact that ethanol decreases ROS levels in catalase-inactivated cells without affecting their T50 values (Fig. 4Go), may be explained because the levels of ROS in the ethanol-treated cells are still above those of control cells, with these levels being sufficient to cause destabilization manifested as a decreased T50 value (Fig. 5Go). Furthermore, in studies on the effects of ethanol on erythrocyte ROS levels, the results clearly showed that ethanol is acting as an antioxidant and not as a pro-oxidant in human erythrocytes.

It therefore appears that ethanol does not induce significant oxidative stress in the human erythrocyte, and these data are in agreement with previous studies (Seeman et al., 1971Go), in which it was found that low ethanol concentrations could protect erythrocytes against haemolysis. Although the mechanism for this protective effect is unknown, it has previously been suggested (Halliwell and Gutteridge, 1999Go) that ethanol can serve as a hydrogen donor in the elimination of the hydroxyl radical with formation of water and the 2-hydroxyethyl radical. The latter is a much less reactive free radical species than hydroxyl radical, and can also be self-eliminated by dimerization to form relatively unreactive 2,3-butanediol that is neither a free radical nor a ROS (Halliwell and Gutteridge, 1999Go).

The studies discussed above lead to the conclusion that, in vitro and possibly in vivo, erythrocyte damage in response to ethanol is not a result of oxidative metabolism of ethanol either inside or outside the erythrocyte but that it is a consequence of either a direct effect of unmetabolized ethanol or the effect of an ethanol metabolite(s) other than acetaldehyde. FAEE are present in the circulatory system following ethanol metabolism to these compounds in a number of cells and tissues, although probably not directly in erythrocytes (Gorski et al., 1996Go). These and other workers (Doyle et al., 1994Go; Saghir et al., 1999Go) have shown that even in alcoholics, the exposure of erythrocytes to FAEE is only transient, and that FAEE are readily degraded by these cells. Our in vitro studies on FAEE have shown that they can be incorporated into erythrocytes and can cause elevated spontaneous haemolysis (up to 26% above control levels in the presence of albumin) and a small but significant increase in membrane PS externalization. Thus, although FAEE do have effects on erythrocytes in vitro, it remains unclear at this stage as to whether there is a significant effect in vivo, as the esters may be eliminated from the erythrocytes by the esterase activity of the cells.

In summary, we conclude that the damage to erythrocytes which occurs on in vitro exposure to ethanol may be caused, at least in part, by unmetabolized ethanol directly, rather than by the oxidation of ethanol to acetaldehyde or its conversion to FAEE. Suggestions that ethanol can directly damage the structures of membranes from erythrocytes and other cell types have previously been made (Trandum et al., 1999Go; Wirkner et al., 1999Go), and a mechanism has been proposed in which ethanol can decrease free radical levels (Halliwell and Gutteridge, 1999Go). Current studies in our laboratory are involved in determining how such effects may occur, and to establish whether FAEE may have an in vivo effect on erythrocytes.


    ACKNOWLEDGEMENTS
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
We thank J. K. Hobbs, D. Jennings and S. M. Polutnik for assistance with gas chromatographic analyses.


    FOOTNOTES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
* Author to whom correspondence should be addressed at: Department of Biomedical Sciences, Irvine Hall, Ohio University, Athens, Ohio 45701, USA. Back


    REFERENCES
 TOP
 FOOTNOTES
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 ACKNOWLEDGEMENTS
 REFERENCES
 
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