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* Laboratoire de Physiologie, et
Laboratoire de Biologie Cellulaire, UFR des Sciences Médicales et Pharmaceutiques, Besançon, France
1To whom correspondence should be addressed. E-mail: alain.berthelot{at}univ-fcomte.fr.
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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KEY WORDS: • primary rat hepatocyte cultures • extracellular magnesium concentration • apoptosis
Magnesium (Mg) possesses a large variety of biological functions, including structural, catalytic and regulatory roles, making Mg deficiency a potential health hazard. In most industrialized countries, Mg intake has decreased over time and is marginal in the entire population (1). Mg deficiency has been shown to lead to an increased incidence of heart diseases, hypertension, total stroke and atherosclerosis [for review, see (2)]. In addition to the unbalanced nutrition, the impaired absorption and increased excretion that occur in certain diseases (e.g., diabetes, alcoholism) and with drug therapy (e.g., diuretics, cisplatin) can favor hypomagnesemia (3).
The involvement of free radicals in tissue injury induced by Mg deficiency causes a reduction of the antioxidant status (4–12) and the accumulation of oxidative products (6–7,10,13, 17) in heart, liver, kidney and skeletal muscle tissues, in RBC and in cultured endothelial and cortical cells. In addition, the prevention of injury induced by Mg deficiency could be achieved through antioxidant treatment (4,6,9). Finally, genes encoding for enzymes involved in cellular protection against oxidative stress have been shown to be up-regulated in thymocytes from Mg-deficient rats (18).
In addition to the well-described induction of necrosis, oxidative damage can also lead to cell death by apoptosis (19–22). The involvement of reactive oxygen species (ROS) and related secondary oxidant species such as lipid hydroperoxides in apoptosis was demonstrated recently. Inducers of apoptosis are able to increase free radicals (23–26); antioxidants and free radical scavengers have been reported to inhibit apoptosis (19,26–30), and finally, ROS per se have been shown to induce apoptosis (28,31,32). A cDNA array analysis of molecular events associated with Mg deficiency in neutrophils revealed an induction of genes encoding proteins involved in apoptosis (33). Other studies have shown a beneficial effect of Mg treatment to decrease apoptosis in neuronal and cardiac cells after hypoxia/ischemia (34–36).
Although the liver plays a major role in Mg homeostasis, as shown by the accumulation or release of very large amounts of Mg upon specific metabolic stimulation (37,38), to our knowledge, only the group of Günther et al. has reported on the susceptibility of liver to Mg deficiency by showing that Mg deficiency enhanced lipid peroxidation in rat liver (16) and in cultured rat hepatocytes (17). The aim of the present study was to examine the effects of Mg on the occurrence of apoptosis in primary cultures of rat hepatocytes. For this purpose, after overnight attachment, rat hepatocytes were maintained for up to 72 h in serum-free media containing low (0 to 0.4 mmol/L) or high (2 and 5.6 mmol/L) concentrations of Mg and compared with 0.8 mmol/L, i.e., the physiological extracellular Mg concentration (14,16,39). Low extracellular Mg concentrations (0–0.4 mmol/L) are associated with moderate and severe human hypomagnesemia [0.01–0.5 mmol/L, (40)], and high concentrations (2 and 5.6 mmol/L) with human hypermagnesemia after therapeutic use [3.5–5 mmol/L, (41)].
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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Collagenase type I, Glutamax Williams’ E medium (+WME), Williams’ E medium without magnesium sulfate (-WME), HBSS, PBS, fetal calf serum (FCS), glutamine, trypsin (0.5 g/L)-EDTA (0.2 g/L) and RNase A were purchased from InVitrogen (Paisley, UK). Hydrocortisone hemisuccinate, insulin, Hoechst 33528 and propidium iodide (PI) were from Sigma Chemical (St. Louis, MO). Magnesium sulfate was from Prolabo (Fontenay, France). All other reagents were of analytical grade.
Animals.
Male Wistar rats weighing 230–270 g were obtained from Charles River (St Germain sur l’Arbresle, France) and housed in metal cages with a 12-h light:dark cycle. All rats had free access to food and filtered tap water.
Isolation and culture of rat hepatocytes.
Rats were anesthetized with thiopental (Pentothal; Abbott, Rungis, France) administered intraperitoneally (100 mg/kg body). Hepatocytes were isolated by a two-step collagenase perfusion, as previously described (42). Cell viability, estimated by Trypan blue exclusion, was >80%. Isolated hepatocytes were resuspended in +WME supplemented with FCS (10%), insulin (4 mg/L), hydrocortisone (10-5 mol/L) and gentamicin (50 mg/L). Cell incubations were performed at 37°C in a humidified atmosphere of 95% air/5% CO2. After 4 h, the medium was renewed to remove unattached hepatocytes and after 12–14 h, cells were shifted to a serum-free medium, consisted of -WME supplemented with glutamine (2 mmol/L), insulin (4 mg/L), hydrocortisone (10-5 mol/L) and gentamicin (50 mg/L). At this time point, cultures were exposed to various concentrations of magnesium sulfate (Mg: 0–5.6 mmol/L). Subsequently, the medium was changed daily.
Determination of intracellular Mg concentration.
Rat hepatocytes were seeded onto home-prepared collagen-coated 24-well plates at a density of 0.2 x 106 cells/well. At the appropriate time points after treatment, hepatocyte monolayers were washed once with ice-cold PBS. Samples were prepared as described by Günther et al. (17), except that cell ash was dissolved in 1% HNO3. The Mg concentration was measured by atomic absorbance spectrophotometry (model Z5000; Hitachi, Tokyo, Japan).
DNA synthesis measurement.
Rat hepatocytes were seeded onto home-prepared collagen-coated 24-well plates at a density of 0.2 x 106 cells/well. At the appropriate time points after treatment, hepatocyte monolayers were labeled with 5-bromo-2'-deoxyuridine (BrdU) and processed as previously described (43). BrdU incorporation was measured using the Cell Proliferation kit (Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s instructions. The absorbance was read at 450 nm against a reference wavelength of 620 nm in a microtiter plate.
Oxidative stress evaluation.
Rat hepatocytes were seeded onto home-prepared collagen-coated six-well plates at a density of 1 x 106 cells/well. At the appropriate time points after treatment, hepatocyte homogenates were prepared as previously described (44) and were frozen at –80°C until analysis. The reduced glutathione (GSH) and the TBARS concentrations were determined as previously described (45).
Protein determination.
The protein concentration was determined by the bicinchoninic acid protein assay kit, according to the manufacturer’s instructions (Sigma Chemical) and bovine serum albumin was used as a standard.
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Apoptosis evaluation |
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TOPABSTRACTMATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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Rat hepatocytes were seeded onto collagen-coated 8-well culture slides at a density of 0.25 x 106 cells/well (Becton Dickinson, Bedford, MA). At the appropriate time points after treatment, hepatocyte monolayers were washed once with PBS, fixed for 10 min with 40 g/L paraformaldehyde in PBS at room temperature, washed twice with PBS and incubated with RNase A (100 mg/L) in fresh +WME for 5 min at room temperature. Cells were then stained for 30 min at 37°C in the dark with Hoechst 33258 (10 mg/L), washed twice with PBS and the slides mounted for fluorescence microscopy. Cells undergoing apoptosis displayed highly fluorescent nuclei, typically pyknotic and with condensed chromatin. Transforming growth factor-ß (TGF-ß) was used as a positive control for apoptosis (43,46).
Flow cytometric analysis.
Rat hepatocytes were seeded onto home-prepared collagen-coated six-well plates at a density of 1 x 106 cells/well. At the appropriate time points after treatment, hepatocyte monolayers were washed once with ice-cold PBS. Cells from one well per group were detached by trypsinization and washed twice with ice-cold PBS. Cells were immediately fixed with ice-cold ethanol (70%) and kept for up to 7 d at 4°C. Ethanol was removed by centrifugation (70 x g for 3 min) and cells were then washed twice in PBS. Cells were resuspended in PBS containing 100 mg/L RNase A at room temperature and stained further in the dark with 10 mg/L PI. Flow cytometric analysis was performed with an ALTRA flow cytometer (Beckman Coulter, Villepinte, France) and with a 610-nm band-pass filter. Each analysis was performed on at least 10,000 events. Data analysis was performed using the expo32 program and each measurement of the DNA content was obtained from the monovariate DNA-PI histogram gated to exclude debris and doublets on the peak/area DNA content histogram. The population of apoptotic cells was obtained from the subdiploid population and expressed as the percentage in the whole cell population. Analysis of freshly isolated rat hepatocytes, which was used to validate the quality of sample preparation, showed a typical ploidy profile, i.e., primarily tetraploid cells with a smaller proportion of diploid and octoploid hepatocytes, as previously reported (47,48).
Determination of caspase-3 activity.
Rat hepatocytes were seeded onto home-prepared collagen-coated six-well plates at a density of 1 x 106 cells/well. At the appropriate time points after treatment, hepatocyte monolayers were washed once with ice-cold PBS. Cells from two wells per group were scraped in PBS and pelleted at 70 x g for 3 min at 4°C. Cell pellets were frozen and stored at -80°C until use. Caspase-3 activity was measured with a commercially available kit (Interchim, Montluçon, France), according to the manufacturer’s instructions. Standard curves were obtained using 7-amino-4-methylcoumarin. Caspase activity was determined using a fluorescence spectrophotometer (excitation at 355 nm and emission at 460 nm).
Statistical analysis.
Statistical comparisons among experimental cultures were performed by one-way (Mg concentration as factor) and two-way (Mg concentration and culture time as factors) ANOVA followed by Tukey’s test. Differences were considered significant at P < 0.05.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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After 72 h of culture in the presence of 0.8 mmol/L Mg, the intracellular Mg concentration of hepatocytes was 77.5 ± 0.5 nmol/mg protein. The intracellular Mg concentration was not different in rat hepatocytes exposed to extracellular Mg below this physiological concentration. In contrast, supplementation of medium with high Mg concentrations, 2 and 5.6 mmol/L Mg, increased the intracellular Mg concentrations 1.6- and 2.7-fold, respectively, compared with medium containing physiological Mg concentrations (P < 0.05).
Hepatocyte morphology and viability.
During the overnight attachment preculture in +WME supplemented with FCS, insulin and hydrocortisone, hepatocytes progressively spread and flattened, until they formed a confluent monolayer as assessed by inverted phase-contrast light microscopy (photomicrographs not shown). The effect of shifting hepatocyte monolayers to serum-free medium containing various concentrations of Mg (0 to 5.6 mmol/L) on cell morphology and overall monolayer integrity was assessed over time (Fig. 1). After 24 h, rounded floating cells were observed in all cultures (photomicrographs not shown). After 48 h of treatment, morphological differences were observed between Mg-treated cultures; these were exacerbated after 72 h of treatment. In Mg-free cultures, rat hepatocytes formed sheets of rounded cells and monolayers had largely shrunk (Fig. 1A). These morphological features were observed in cultures treated with 0.2 and 0.4 mmol/L Mg, although to a lesser extent than in Mg-free cultures (Fig. 1B,C). In cultures treated with 0.8 mmol/L Mg, the major feature was the presence of numerous rounded floating cells. Small foci without adherent cells were occasionally observed on cell monolayers (Fig. 1D). In contrast, rat hepatocytes treated with 2 and 5.6 mmol/L Mg spread out as a confluent monolayer, despite the presence of a few rounded cells (Fig. 1E, F).
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After 72 h of treatment, DNA synthesis, as assessed by BrdU incorporation, was not affected by the exposure of rat primary hepatocyte cultures to 0–5.6 mmol/L Mg (data not shown).
Rates of oxidative stress.
After 72h of treatment, GSH concentration was significantly lower in Mg-free monolayers compared with the other conditions, as well as in cell monolayers cultured in the presence of 0.2 mmol/L Mg compared with 0.8 mmol/L Mg (Table 1). Similarly, lipid peroxidation, as assessed by the measurement of TBARS concentration, was significantly higher in Mg-free monolayers compared with cultures exposed to 0.2–5.6 mmol/L Mg, and in cultures containing 0.2 and 0.4 mmol/L Mg compared with those containing 0.8–5.6 mmol/L Mg (Table 1).
Apoptosis.
Hepatocellular apoptosis was characterized qualitatively by fluorescence microscopy using the DNA-binding fluorochrome Hoechst 33258. No major differences appeared between cultures after 24 h of culture in the presence of the various concentrations of Mg (photomicrographs not shown). In contrast, after 48 and 72 h of culture, nuclei that displayed the typical figures of apoptotic morphology appeared more numerous in Mg-free cultures compared with nuclei of cells cultured with 0.8 or 5.6 mmol/L Mg (Fig. 2). Cultures treated with TGF-ß, showed considerable chromatin condensation and nuclear fragmentation (Fig. 2A). Similar, although less pronounced changes, were observed in Mg-free cultures (Fig. 2B). Compared with Mg-free cultures, most hepatocytes from cultures containing 0.8 mmol/L or 5.6 mmol/L Mg displayed normal chromatin and nuclear structure, and their nuclei were weakly stained (Fig. 2C,D).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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Extracellular Mg deficiency was not associated with corresponding decreases in intracellular Mg concentrations in primary cultures of rat hepatocytes. In accordance with our results, Günther et al. (5,17) showed that total Mg concentration in rat liver and in cultured rat hepatocytes was not significantly affected by decreases in extracellular Mg concentration. Fagan et al. (38) showed that changes in cellular Mg concentration occur only under conditions of hormonal stimulation in rat hepatocytes. This is in contrast with observations in other organs (5,49). However, we found that supplementation with high extracellular Mg concentrations increased cellular Mg, in agreement with results obtained in endothelial cells (50). This leads to the conclusion that the deleterious effect of Mg deficiency on rat hepatocytes in primary cultures is due to an extracellular lack of Mg.
In accordance with previous publications, which have shown that Mg deficiency enhanced lipid peroxidation in both rat livers (10,14,16) and cultured rat hepatocytes (17), we found that lipid peroxidation was significantly enhanced in cultures with low extracellular concentrations of Mg (0 to 0.4 mmol/L). Several authors have hypothesized that this might be due in part to changes in Na+ and Ca2+ homeostasis. Indeed Mg deficiency leads to an accumulation of Na+ and Ca2+ in RBC (51). Mitochondrial respiratory activity, which is regulated by Ca2+, might be affected, leading to ROS production during Mg deficiency (9,51). Nevertheless, other authors have shown that K+, Na+ and Ca2+ concentrations in rat livers were essentially unaffected by dietary Mg deficiency compared with other organs (5,16). These authors suggested that enhanced lipid peroxidation occurring in livers of Mg-deficient rats was related to the presence of large amounts of iron in liver, higher than in other tissues (5,16) because Fe could substitute for Mg at the intra- and extracellular side of the cell membrane and thereby initiate radical chain reactions via the Fenton reaction. Addition of a specific Fe chelator to rat hepatocyte cultures treated with Mg concentrations below the physiological concentration completely inhibited lipid peroxidation (17), indicating that enhanced lipid peroxidation during Mg deficiency was indeed due to Fe.
Also, in agreement with previous observations of Rimbach et al. (10) showing that an impaired Mg bioavailability in rats was accompanied by a decrease in liver glutathione concentration, we have shown in this study that GSH concentration was significantly decreased in primary rat hepatocyte cultures treated with low Mg concentrations (0 and 0.2 mmol/L). Indeed, the involvement of free radicals in Mg deficiency is well documented in vivo in animals and in vitro in cell cultures; many of these publications provided direct support for a reduction in antioxidant content, i.e., a decrease in GSH concentration in heart and liver, in RBC and in cultured endothelial and cortical cells (4,6,8–12), and a reduction in liver ascorbate synthesis (52) and vitamin E concentration (5).
In the present study, we clearly showed that the apoptosis rate was increased in rat hepatocyte cultures in the presence of low concentrations of extracellular Mg (0 and 0.2 mmol/L). Apoptosis was assessed by the combined detection of morphological changes using a DNA-binding fluorochrome (30,43), the quantification of subdiploid population using analysis of DNA content by flow cytometry (53) and the quantification of caspase-3 activity because caspase-3 plays an important role in the effector phase of apoptosis (54). This selection of apoptotic markers was made according to the recommendations of Gomez-Lechon et al. (55), who identified these sensitive markers in cultured rat hepatocytes.
Because the involvement of ROS and related secondary oxidant species in apoptosis has been shown recently (19,23–32), it suggests that Mg deficiency–induced ROS production, as assessed by GSH depletion, could be related to apoptosis. A recent study revealed that in neutrophils, Mg deficiency induced genes encoding proteins involved in stress and apoptosis (33). Indeed, these authors obtained an increase of HSP 70 (83-fold increase), caspase-3 (9-fold increase), bad (46-fold increase) and p21-activated kinase (8-fold increase) in Mg-deficient rats compared with control rats. Other studies have shown an indirect role of Mg in apoptosis because Mg treatment can reduce apoptosis in neuronal and cardiac cells after hypoxia/ischemia (34–36).
The increased apoptosis rate in primary cultures of rat hepatocytes in the presence of low extracellular Mg concentration might be due in part to loss or impairment of cell-matrix interactions. Indeed several authors reported that cell adhesion is Mg-dependent for different cell types (56–58). For liver, Ponce et al. (59) showed that hepatocyte adhesion to laminin was Mg dependent. More recently, Menon et al. (60) showed that attachment of hepatocytes to collagen IV and laminin was promoted by Ca2+ and Mg2+. However, other important proteins are present in the liver extracellular matrix, such as fibronectin, collagen type I, III, V, VI as well as various proteoglycans (61). Further studies are thus required to evaluate the modulation of hepatocyte adhesion to extracellular matrix proteins by Mg, especially for hepatocytes already attached to culture support, and especially concerning the finding that Mg concentrations, which have been shown to be maximal for hepatocyte adhesion to laminin, were far lower (0.08 mmol/L) than the 0.2 mmol/L extracellular Mg concentration for which we observed a significant increase in apoptosis rate compared with cultures in the presence of the physiological 0.8 mmol/L Mg concentration. It is thus unlikely that impairment of adhesion of hepatocytes was the reason for cell detachment in culture conditions with low Mg concentrations.
Because cell viability is the result of a balance between apoptosis and proliferation, we evaluated the BrdU incorporation into cultured hepatocytes treated with various extracellular Mg concentrations for up to 72 h. We showed that increased extracellular Mg did not stimulate DNA synthesis in our cultures. This is in contrast to other publications, showing that extracellular Mg acts as a positive modulator of cell proliferation in normal keratinocytes (62), endothelial cells (63) and neural cells (64). This indicates that Mg has no effect on cell proliferation in primary cultured hepatocytes, which are considered almost nonproliferative cells in culture (65,66).
The results of the present study have not clearly identified a beneficial effect of high (2 and 5.6 mmol/L) concentrations of Mg on hepatocellular viability. At these concentrations, the well-documented spontaneous apoptosis occurring in rat hepatocyte cultures over time (67,68) and observed in this study in the presence of physiological concentrations of Mg could not be counteracted.
In sum, the present study shows that extracellular Mg deficiency induces apoptosis in primary cultures of rat hepatocytes, most probably by an oxidative stress–related mechanism. This hypothesis is strengthened by preliminary data showing that, as expected (69), a N-acetylcysteine supplementation of Mg-free medium restored the GSH concentration (a 12-fold increase compared with nonsupplemented Mg-free medium), and this was associated with reduced activation of caspase-3 (a 65% decrease compared with nonsupplemented Mg-free medium), and a decreased lipid peroxidation rate (a 75% decrease compared with nonsupplemented Mg-free medium). Further studies are currently underway to clarify the relationship between Mg deficiency and the decrease in GSH that subsequently induces increases in lipid peroxidation.
In conclusion, our study shows that extracellular Mg deficiency has a negative effect on the survival of cultured rat hepatocytes by inducing apoptosis; however, supplementation of extracellular Mg did not reduce spontaneous apoptosis occurring over time in rat hepatocyte cultures.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Manuscript received 7 March 2003. Initial review completed 24 March 2003. Revision accepted 1 May 2003.
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LITERATURE CITED |
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TOPABSTRACTMATERIALS AND METHODSApoptosis evaluationRESULTSDISCUSSIONLITERATURE CITED |
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