Muscle Research Group, Institut d'Investigacions Biomèdiques August Pi i Sunyer, Unitat de Bioquímica and Department de Medicina, Universitat de Barcelona, Spain
Received 6 September 1999; in revised form 19 January 2000; accepted 26 January 2000
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ABSTRACT |
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INTRODUCTION |
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In mammals, chronic ingestion of EtOH by pregnant females may produce serious damage to the fetus, and the worst consequences may be to the nervous system (Schenker et al., 1990). Guerri et al. (1990) demonstrated that EtOH inhibited DNA and RNA synthesis and decreased DNA and protein concentrations in primary cell cultures of astrocytes. These effects were more drastic when the astrocytes had been obtained from embryos of alcoholic rats. Davies and Cox (1991) showed that the apparent effects of EtOH on astrocyte differentiation were a consequence of the effects of EtOH on cell proliferation. It has been proven that the effect of EtOH on the nervous system depends on the time of exposure, since the expression of growth factors and receptors, which are believed to interact with EtOH, changes during proliferation and differentiation (Brodie and Vernadakis, 1992
).
EtOH modifies the hepatic regeneration of hepatectomized rats (Wands et al., 1979) and is considered to induce neoplasia in the digestive system (Doll and Peto, 1981
). Similarly EtOH-mediated effects have also been described on the proliferation of primary rat hepatocyte cultures (Carter and Wands, 1988
) and various mammalian cell lines, including BALB/c mouse 3T3 fibroblast (Resnicoff et al., 1993
), HL-60 human myeloid leukaemia cells (Cook et al., 1990
; Datta et al., 1990
), human bone cells (Friday and Howard, 1991
), human T cells (Imperia et al., 1984
), and rat hepatic tumour cells (Higgins and Borenfreund, 1986
; Higgins, 1987
). Although the acute effect of EtOH on calcium transients from cultured human myotubes has been studied (Nicolas et al., 1998
), no data have been produced which analyse the effects of EtOH on skeletal muscle proliferation or differentiation.
The aim of this study was to analyse the direct effects of EtOH on muscle cell proliferation and differentiation as well as its effect on protein and DNA levels by using a primary cell culture model derived from muscular satellite cells, which closely resembles the maturation process of muscle in vivo in the absence of any other tissue.
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MATERIALS AND METHODS |
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Cell cultures
Primary cultures were obtained as described by Gautron (1988) with some modification. Each primary culture preparation was obtained by seeding the satellite cells obtained from the posterior limbs of two rats (SpragueDawley) of 8 to 12 weeks of age. After killing the animals by ether anaesthesia, the skin and the conjunctive tissue of the limbs were removed. Each muscle was then carefully dissected to discard fat and connective tissues and washed several times in sterile phosphate buffered saline (PBS) (pH 7.4). The muscles were then minced with scissors and digested for 2 h in 40 ml of 0.17% pronase diluted in Ham's F-12 medium (Gibco Laboratories, Grand Island, NY, USA) pH 7.35, plus 10% fetal calf serum (FCS) (Gibco Laboratories) at 37°C. The satellite cells obtained from the digested muscles were collected in the supernatant after centrifugation of the digested muscles in pronase solution (90 g for 3 min). This supernatant was centrifuged (350 g for 20 min) and the satellite cells in the pellet were resuspended in Dulbecco's modified Eagle's medium (DMEM) (Gibco Laboratories) supplemented with 10% FCS and 10% horse serum (HS). Cells were resuspended to a final concentration of 10 000 cells/ml and distributed in flasks (60 000 cells/25 cm2 flask, Costar, Cambridge, MA, USA). To facilitate adhesion of muscular cells to the plastic surface, and to avoid attachment of fibroblasts, flasks were precoated with gelatine (0.5% in water). Normally, we obtained about 1 million cells (~15 to 20 flasks of cells)/primary culture preparation. The cells were grown in DMEM plus 10% FCS and 10% HS supplemented with penicillin (0.1 mg/ml) and streptomycin (100 U/ml) (Gibco Laboratories) at 37°C in a humidified atmosphere of 5% CO2.
Subcultures were generated to obtain a suitable number of cells for the cell proliferation assays. Four days after seeding, the cells were still proliferating and had to be split 1:4 to avoid differentiation (see below) and induce further proliferation. After removing the medium from the flasks, cells were washed in PBS and then incubated with 1 ml of trypsin (0.25% in PBS) at 37°C for 3 min. Once the cells were detached, addition of fresh complete medium stopped the trypsin activity (Sigma Chemical Co., St Louis, MO, USA). The cells were then reseeded as described above. This tissue culture model, or similar (using embryonic myoblasts, instead of satellite cells) is widely used for various purposes, including the study of myogenesis.
To obtain primary cell cultures, satellite cells were isolated enzymatically from rat skeletal muscle and grown in culture. These cultures last about 3 weeks. During the first 45 days, cells proliferated and then started to fuse and differentiated into adult multi-nucleated skeletal muscle cells. All the morphological changes were monitored under a phase contrast microscope (Fig. 1. ). Differentiation was most evident during the second week of culture and, concomitant with morphological changes, a series of biochemical phenomena also occurred, such as changes of the isozyme pattern of creatine kinase (CK), from the embryonic form (BB) to muscle adult form (MM). Finally cells start to degenerate and die (Alterio et al., 1990
). The tissue culture was differentiating when cells started to fuse to form myosacs and finally myotubules. This process was also reflected in the changes in the percentages of each isoenzyme of the skeletal muscle enzyme CK: from 31.8% (MM), 14.8% (MB), and 53.5% (BB), on day 5, to 61% (MM), 26.7% (MB), and 12.7% (BB) on day 12. When these values were compared to those of skeletal muscle in vivo, the tissue culture model did not show the same degree of differentiation because of the lack of innervation.
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Effect of EtOH on cell proliferation
The effects of EtOH on cell proliferation were studied during the first 5 days of the subculture. On day 2 of the subculture, EtOH was added to the medium at the following concentrations, 10 mM, 25 mM, and 100 mM. The EtOH concentrations chosen were within the range usually found in human plasma of chronic alcoholics (up to 100 mM EtOH) (Cussó et al., 1989). To evaluate changes in proliferation rates, cells were grown in the presence of 2.4 nM [3H]thymidine (0.2 µCi/ml) (Dupont, Boston, MA, USA) for 18 h (Cleaver and Holford, 1965
). Cells were then washed three times in PBS and lysed with 0.1 M NaOH (0.5 ml/flask). The radioactivity incorporated was measured in three aliquots of lysate (50 µl) in a ß-scintillation counter. The value was referred to the amount of protein. In each experiment, at least two flasks were used for each EtOH concentration and time of assay (1 to 4 days).
Effect of EtOH on cell differentiation
These assays were started by adding EtOH to the cells on day 5 of culture, when the cells stopped dividing. We measured the percentage of each of the three CK isozymes, from day 6 to day 12 of culture, which spans the differentiation programme. In each experiment, at least two different concentrations of EtOH were tested. Total CK activity was assayed as described by Oliver (1955). The amount of each CK isozyme was measured by the electrophoresis Cardiotrac-CK kit (Corning, Cambridge, MA, USA), based on the method described by Gerhardt and Waldenstrom (1979). In brief, the three isozymes (MM, MB, and BB) were separated electrophoretically in agarose gels and then detected by the fluorescence produced by NADH, the final product of a series of reactions. The fluorescence was measured immediately in a densitometer.
Cell extracts, protein, and DNA measurements
Cell extracts were obtained by scraping the cells into 0.5 ml of buffer (50 mM HClTris pH 7.2, 4 mM EDTA, and 30 mM ß-mercaptoethanol); the flask and the suspension was sonicated with a Branson sonifier (60 W, 1 s x 5). Lysates were clarified by centrifugation (12 000 rpm for 10 min, at 4°C) and the enzymatic assays were performed in the supernatant as described above. Protein was measured following Bradford (1976) using Bio-Rad reagent (Bio-Rad Laboratories, Hercules, CA, USA) and a standard of bovine serum albumin (0.524 µg). DNA was determined following Labarca and Paigen (1980), where the Hoechst 33258 reagent {bisbenzimide: [(2-[2- (4-hydroxyphenol)-6-benzimidazolil-6 (1-methyl-4-piperazil) benzimidazol]} (Sigma Chemical Co.) produces fluorescence when bound to double-stranded DNA.
Statistics
Statistical comparisons of the effect of various EtOH concentrations on different culture days were performed by analysis of variance (one- or two-way ANOVA). Variations between group means were considered significant at P < 0.05. Those variations were evaluated by the TukeyKramer multiple comparison test. Variations were considered significant at P < 0.05.
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RESULTS |
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DISCUSSION |
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Our results are in agreement with other studies reported by several groups, which consider that the more undifferentiated the cells, the more sensitive they are to the action of EtOH (Guerri et al., 1990; Miller, 1992
). However, after being exposed to the action of EtOH, skeletal muscle cells seemed to become adapted to its presence and there was little inhibition of cell proliferation.
The effect of EtOH on cell differentiation has been described in several cellular models, mostly from the nervous system (Davies and Vernadakis, 1986; Davies and Cox, 1991
). In our experimental model, the action of EtOH on cell differentiation does not seem to be dose dependent, as it occurred in the proliferative phase. Nevertheless, despite the presence of EtOH, cells reached a very similar degree of differentiation, as judged by morphological observations (data not shown) and by the pattern of CK isozymes. This implies that, although the action of EtOH can slow down the differentiation programme, it cannot block eventual differentiation. Thus, taken together, our results suggest that EtOH inhibits skeletal muscle proliferation and delays its differentiation, although the mechanisms remain to be determined.
If we consider that EtOH is scarcely metabolized in skeletal muscle, these effects cannot be the consequence of the action of the products of the EtOH degradation. The changes in cell proliferation and differentiation may be due to the action of the EtOH molecules on cell structures. It has been shown that EtOH can physically interact with cell membranes, disorganizing them and, thus, modifying the processes occurring in these cellular structures. Among such processes, it has been demonstrated that EtOH perturbs the adenylyl cyclase system (Hoek and Rubin, 1990). This system is important for both the proliferation and the differentiation processes, given that several proteins that participate in these processes are phosphorylated by the adenylate cyclase system (Pennington, 1988
). The effect of EtOH on the processes depending on Ca2+, which acts as both signal transducer and in the Ca2+ ATPase system, can also be affected by EtOH action on cell membranes (Rubin et al., 1976
; Hoek et al., 1992
). Finally, the interaction of EtOH with cell membranes makes them less sensitive to factors that potentiate cell proliferation and differentiation (Pennington et al., 1985
; Sonntag and Boyd, 1988
; Brodie and Vernadakis, 1992
; Resnicoff et al., 1993
).
Finally the studies on the effect of EtOH on protein content seem to demonstrate that this toxin has no effect on its levels. However, we cannot rule out the possibility that EtOH has an effect on muscle protein turnover in vivo (Preedy and Peters, 1990), a phenomenon that appears after years of EtOH consumption.
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ACKNOWLEDGEMENTS |
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FOOTNOTES |
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1 Present address: Fels Institute for Cancer Research and Molecular Biology, Temple University School of Medicine, 3307 N Broad Street, Philadelphia, PA 19140, USA.
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