Several studies support the concept of a diabetic cardiomyopathy in the absence of discernible coronary artery disease, although its mechanism remains poorly understood. We investigated the role of glucose and palmitic acid on cardiomyocyte apoptosis and on the organization of the contractile apparatus. Exposure of adult rat cardiomyocytes for 18 h to palmitic acid (0.25 and 0.5 mmol/l) resulted in a significant increase of apoptotic cells, whereas increasing glucose concentration to 33.3 mmol/l for up to 8 days had no influence on the apoptosis rate. However, both palmitic acid and elevated glucose concentration alone or in combination had a dramatic destructive effect on the myofibrillar apparatus. The membrane-permeable C2-ceramide but not the metabolically inactive C2-dihydroceramide enhanced apoptosis of cardiomyocytes by 50%, accompanied by detrimental effects on the myofibrils. The palmitic acid–induced effects were impaired by fumonisin B1, an inhibitor of ceramide synthase. Sphingomyelinase, which activates the catabolic pathway of ceramide by metabolizing sphingomyeline to ceramide, did not adversely affect cardiomyocytes. Palmitic acid–induced apoptosis was accompanied by release of cytochrome c from the mitochondria. Aminoguanidine did not prevent glucose-induced myofibrillar degeneration, suggesting that formation of nitric oxide and/or advanced glycation end products play no major role. Taken together, these results suggest that in adult rat cardiac cells, palmitic acid induces apoptosis via de novo ceramide formation and activation of the apoptotic mitochondrial pathway. Conversely, glucose has no influence on adult cardiomyocyte apoptosis. However, both cell nutrients promote degeneration of myofibrils. Thus, gluco- and lipotoxicity may play a central role in the development of diabetic cardiomyopathy.

It has long been known that diabetes has major cardiovascular effects and is a risk factor for ischemic heart disease. Moreover, patients with diabetes have a markedly adverse course after myocardial infarction, with high rates of postinfarction heart failure and death (1,2). Several studies support the concept of a specific diabetic cardiomyopathy, i.e., cardiac abnormalities in the absence of discernible coronary artery disease, hypertension, or valvular disease (3,4). Several mechanisms for the pathogenesis of diabetic cardiomyopathy, such as endothelial dysfunction, autonomic dysfunction, metabolic derangement, and interstitial fibrosis, have been proposed. Furthermore, abnormalities in the contractile proteins could be responsible for the mechanical defects in the heart of patients with diabetes. For example, shifts in cardiac myosin heavy chain probably contribute to the impaired cardiac function in the heart of chronic diabetic rats (5). However, the precise causative factors that lead to diabetic cardiomyopathy remain to be investigated.

Both cell nutrients glucose and free fatty acids are pathologically elevated in the circulating blood of diabetic patients and have been suggested to have adverse effects on cell function, so called “glucotoxicity” and “lipotoxicity” (6,7). Loss of myofibrils is the most obvious structural change in dilated cardiomyopathy (8), and sarcomeric disarray is characteristic of failing hearts (9,10). However, the possible role of glucose and fatty acids on myofibrillar organization has not been addressed.

Apoptosis of cardiomyocytes was shown recently to play a central role in the development of heart failure (11). Saturated fatty acid–induced apoptosis has been demonstrated in neonatal rat cardiomyocytes (12,13,14). However, whether fatty acids also can induce apoptosis in adult cardiomyocytes, which are postmitotic, and the underlying mechanism remain to be investigated.

Depending on culture conditions, cell type, and genetic background, glucose proved to be pro- or antiapoptotic (15,16,17,18,19,20). In neonatal rat cardiomyocytes, serum and glucose deprivation induced apoptosis (15,19), whereas elevated glucose concentrations prevented the cells from hypoxia-induced apoptosis (20). However, the possible role of glucose in adult cardiomyocyte apoptosis has not been investigated in these studies.

The aim of the present study was to investigate the role of palmitic acid (C16:0) and of elevated glucose concentrations on adult cardiomyocyte apoptosis. In addition, the involvement of the ceramide and of the apoptotic mitochondrial pathway was studied. Furthermore, the role of glucose and palmitic acid on the organization of the contractile apparatus was assessed.

Cell culture.

Ventricular cardiac muscle cells of adult female rats (Sprague-Dawley-Ivanovas, 2 months old) were isolated by retrograde perfusion of hearts with collagenase type 2 (Worthington Biochemical, Freehold, NJ) as described (21,22). After perfusion, the heart tissue was minced and incubated at 37°C for another 10 min in KB medium (23) containing collagenase. Cells were cultured in dishes coated with 0.1% gelatin in M-199 supplemented with 20% fetal calf serum (FCS) (Sigma Chemical, St. Louis, MO), 1% penicillin/streptomycin, and 20 mmol/l creatine. For inhibiting growth of contaminating cells, 10 μmol/l 1-β-d-arabinofuranosyl-cytosine was added throughout the culture period. The 20% FCS medium was changed to medium containing 10% FCS after 2, 7, and, for prolonged experiments, 9 days. Basic medium (5.5 mmol/l glucose) was supplemented with 27.8 mmol/l glucose or with fatty acids as described below. Palmitic and oleic acids (Sigma) were dissolved at 10 mmol/l in M-199 medium containing 11% fatty acid–free bovine serum albumin (BSA) (Sigma) under N2 atmosphere, shaken overnight at 37°C, sonicated for 15 min, and filtrated under sterile conditions (stock solution). For control incubations, 11% BSA was prepared, as described above. The effective free fatty acid concentration was determined before and after sterile filtration with a commercially available kit (Wako Chemicals, Neuss, Germany). A control experiment was performed with palmitic acid dissolved first in 10 mol/l NaOH, then diluted to 12 mmol/l in M-199 medium containing 12.5% fatty acid–free BSA (Sigma) under N2 atmosphere, and shaken overnight at 50°C. The resulting free fatty acid concentration and the apoptotic effect were similar (not shown). The calculated concentration of not albumin-bound free fatty acid was ∼0.7 μmol/l for a final concentration of 0.5 mmol/l palmitic acid (24). In some experiments, cardiomyocytes were cultured with 0.5 mmol/l aminoguanidine (Aldrich Chemical Company, Milwaukee, WI), 50 μmol/l sphingomyelinase (Sigma), 15 μmol/l C2-ceramide (Biomol, Plymouth Meeting, PA), 15 μmol/l C2-dihydroceramide (Biomol), or 15 μmol/l fumonisin B1 (Sigma). C2-ceramide, C2-dihydroceramide, and fumonisin B1 first were dissolved in prewarmed 37°C DMSO (Fluka, Buchs, Switzerland) at 5 mmol/l.

Detection of apoptotic cardiac cells and cytochemistry.

The free 3-OH strand breaks resulting from DNA degradation were detected by the terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) technique (25). After washing with phosphate-buffered saline, cultured cardiomyocytes were fixed in 3% paraformaldehyde (20 min at room temperature) followed by permeabilization with 0.2% triton X-100 (15 min at room temperature) and by the TUNEL assay, performed according to the manufacturer’s instructions (In Situ Cell Death Detection Kit, AP; Boehringer Mannheim, Mannheim, Germany). The preparations then were rinsed with Tris-buffered saline and incubated (12 min at room temperature) with 5-bromo-4-chloro-indolyl phosphate/nitro blue tetrazolium liquid substrate system (Sigma). Thereafter, cells were incubated for 1 h at room temperature with a mouse antibody anti-myomesin, a sarcomeric M-line protein (26), followed by a 30-min incubation with fluorescein-conjugated goat anti-mouse (Jackson Immuno Research Laboratories, West Grove, PE) and with rhodamine-phalloidin for staining of F-actin (Molecular Probes, Eugene, OR).

The TUNEL assay detects DNA fragmentation associated with both apoptotic and necrotic cell death; therefore, cardiomyocytes also were treated with a fluorescent annexin V probe (Annexin-V-FLUOS staining kit; Boehringer Mannheim) according to the manufacturer’s instructions. Double staining of cells with propidium iodide and annexin V allows differentiation of apoptotic from necrotic cells.

Mitochondria were stained with anti–cytochrome c and with anti-adenine nucleotide translocator for localization of the mitochondria. Cells were fixed and permeabilized as described above, incubated for 2 h at room temperature with mouse anti–cytochrome c monoclonal antibody (PharMingen, San Diego, CA) and with anti-adenine nucleotide translocator antibody (provided by T. Wallimann, Institute of Cell Biology, Swiss Federal Institute of Technology, Zurich, Switzerland [27]), followed by 1-h incubation with fluorescein-conjugated goat anti-mouse antibody and with Cy3-conjugated donkey anti-rabbit antibody (Jackson Immuno Research). After staining, cells were embedded in Dako fluorescent mounting medium (Dako, Glostrup, Denmark).

Microscopy.

Light and immunofluorescence microscopy was performed with a Zeiss fluorescence microscope (microscope Axiolab, Carl Zeiss, Oberkochen, Germany). Images were produced by a Leica inverted microscope DM IRB/E, a Leica true confocal scanner TCS NT, and a Silicon Graphics workstation. The images were recorded using a Leica PL APO 63× oil or a PL APO 40× oil immersion objective. The system was equipped with an argon/krypton mixed gas laser. Image processing was performed on a Silicon Graphics workstation using Imaris (Bitplane AG, Zurich, Switzerland) 3D multichannel image processing software specialized for confocal microscopy images.

The quality of the myofibrillar structures was quantified by scoring the cells according to the myofibrillar organization (myofibrils aligned in line with stress-fiber–like structures or not), to the sarcomeric disarray (sarcomers parallel to each other or not) and to the extent of myofibrillar structures (estimation of the percentage of the cell filled with myofibrils). Only cells that stained negative in the TUNEL assay were counted.

Mitochondrial isolation.

For analysis of cardiomyocyte mitochondria, cells were washed in phosphate-buffered saline and scraped off into 35 μl of ice-cold buffer containing 20 mmol/l HEPES-KOH (pH 7.4), 10 mmol/l KCl, 1.5 mmol/l MgCl2, 1 mmol/l Na-EDTA, 1 mmol/l EGTA, 1 mmol/l dithiothreitol, 1 mmol/l phenylmethylsulfonyl fluoride, 10 μg/ml leupeptin, and 250 mmol/l sucrose (28,29). Unlysed cells and nuclei were pelleted by 10 min of centrifugation (750g, 4°C). The supernatant was centrifuged at 10,000g for 15 min at 4°C. This pellet, which represented the mitochondrial fraction, then was resuspended in 10 μl of the above described buffer and frozen at −80°C until used.

Western blot analysis.

Mitochondrial fractions were diluted 1:1.5 in SDS–Laemmli buffer (30) and boiled for 5 min. Equivalent amounts of each treatment group were run on 15% SDS polyacrylamide gels. Proteins were transferred electrically to nitrocellulose filters and incubated with a mouse anti–cytochrome c monoclonal antibody (PharMingen; 1 h at room temperature) followed by incubation with horseradish peroxidase–linked anti-mouse IgG (Santa Cruz Biotechnology, Santa Cruz, CA; 1 h at room temperature). After Lumiglo reagent (Phototope-HRP Western blot Detection Kit; Biolabs, Beverly, MA) was added, the emitted light was captured on X-ray film. As a marker, a biotinylated protein molecular weight standard (Biolabs) was run in parallel according to the manufacturer’s instructions.

For adenine nucleotide translocator analysis, the nitrocellulose membrane was stripped for 30 min at 50°C in 40 ml of a watery solution containing 280 μl of β-mercaptoethanol, 5 ml of 0.5 mol/l Tris-HCl (pH 6.8), and 10% SDS, washed for 1 h in Tris-buffered saline containing 0.1% Tween-20, incubated with an anti-adenine nucleotide translocator antibody (see above) followed by incubation with horseradish peroxidase–linked anti-rabbit IgG (Santa Cruz; 1:5,000, 1 h at room temperature), and detected as described above.

Statistical analysis.

Data are presented as mean ± SE and analyzed by analysis of variance with a Bonferroni correction for multiple group comparisons. Cultures were evaluated in a randomized manner by a single investigator (M.Y.D.), who was blinded to the treatment conditions.

Effects of palmitic and oleic acids and of glucose on cardiomyocyte apoptosis.

Exposure of adult rat cardiomyocytes in long-term culture to 0.5 mmol/l palmitic acid for 18 h resulted in an increased number of cardiac cells with TUNEL-positive nuclei (Figs. 1A and C). The increase was 1.5-, 2.1-, and 2.3-fold after 6, 10, and 18 h of treatment, respectively, as compared with control (Fig. 2A). Palmitic acid–induced DNA fragmentation was already detectable at a concentration of 0.1 mmol/l, although only a concentration of 0.25 mmol/l or higher was statistically significant (Fig. 2B). In contrast, oleic acid (0.1–0.5 mmol/l) for 18 h and elevated glucose concentrations (33.3 mmol/l) for up to 8 days did not induce DNA fragmentation (Figs. 1E and 2). When added together with palmitic acid, oleic acid but not glucose reduced its effect on cell death (Fig. 2).

In parallel with the TUNEL assay, treated cardiac cells were incubated with annexin V and propidium iodide to discriminate apoptotic from necrotic cells (Fig. 3). Exposure of cultured cardiomyocytes to palmitic acid for 18 h markedly increased the number of cells that exhibited phosphatidylserine molecules translocated to the outer leaflet of the plasma membrane as revealed by annexin V binding (13.5 ± 2.34% Annexin-V-FLUOS–positive cells in control and 61.8 ± 6.88% in 0.5 mmol/l palmitic acid–treated cardiomyocytes; P < 0.01). Part of these cells had defective plasma membranes, permeable to the DNA-binding dye propidium iodide (7.9 ± 2.77% propidium iodide–positive cells in control and 32.4 ± 4.48% in 0.5 mmol/l palmitic acid–treated cardiomyocytes; P < 0.01). Therefore, the palmitic acid–induced DNA fragmentation, as determined by the TUNEL assay, represents both apoptotic cell death positive for Annexin-V-FLUOS only (29%) and necrotic cell death or late apoptotic stages positive for Annexin-V-FLUOS as well as propidium iodide (32%).

Degeneration of myofibrils upon exposure to palmitic acid or glucose.

Exposure of adult rat cardiomyocytes to palmitic acid for 18 h destroyed both the contractile elements and the cytoskeleton (Figs. 1B and D and 4). The deleterious effect of palmitic acid on myofibrils preceded DNA fragmentation and was already detectable 3 h after administration of palmitic acid (Fig. 5). A short-term increase in the ambient glucose concentration to 33.3 mmol/l for up to 1 day had no effect on the contractile apparatus (Figs. 4 and 6), whereas long-term culture of cardiomyocytes in 33.3 mmol/l glucose for 4 and 8 days induced loss of myofibrillar organization and sarcomeric disarray (Figs. 1F and 6). Degeneration of myofibrils also was observed when the cells were exposed for 4 days to high glucose concentrations added after 9 days in culture, when myofibril formation is mostly completed (Fig. 6). Aminoguanidine (0.5 mmol/l), an inhibitor of nitric oxide synthase and of advanced glycation end product formation (31,32), failed to prevent the deleterious effects of glucose (Fig. 7).

Role of ceramide signaling in palmitic acid–induced cardiac-cell apoptosis and alteration of myofibrillar expression.

Cardiomyocytes cultured with 15 μmol/l of the membrane-permeable C2-ceramide displayed an increase in DNA fragmentation and disturbed contractile elements, mimicking the effects observed in palmitic acid–treated cells (Figs. 8 and 9). This was not observed with the metabolically inactive C2-dihydroceramide, which had no effect on the cells. However, addition of the ceramide synthase inhibitor fumonisin B1 (15 μmol/l) to the culture medium containing 0.5 mmol/l palmitic acid prevented the effects of palmitic acid on cardiomyocyte apoptosis and myofibrillar organization. Sphingomyelinase, which activates the catabolic pathway of ceramide by metabolizing sphingomyeline to ceramide, had no adverse effects on the cells. Increasing the ambient glucose concentration from 5.5 to 33.3 mmol/l had no effect on the modulation of the ceramide pathway.

Palmitic acid–induced cytochrome c release.

Exposure of cardiomyocytes to palmitic acid time-dependently decreased mitochondrial cytochrome c content as compared with control cells (Fig. 10). In contrast, adenine nucleotide translocator expression remained unchanged in the mitochondrial fraction of palmitic acid–treated cells, confirming the integrity of the mitochondrial preparations.

This study shows that palmitic acid induces apoptosis in adult rat cardiomyocytes. Cell death induced by free fatty acid has been postulated to be mediated via formation of ceramide as ceramide is synthesized from long-chain fatty acids (33). In the present study, the cell-permeable ceramide analog C2-ceramide mimicked the deleterious effects of palmitic acid, which were blocked by the ceramide synthase inhibitor fumonisin B1. Sphingomyelinase, which activates the catabolic pathway of ceramide by metabolizing sphingomyeline to ceramide, had no adverse effects on the cardiomyocytes. Thus, de novo ceramide generation is part of the fatty acid–induced cell death in cardiomyocytes. However, the deleterious effects of palmitic acid were not completely blocked by fumonisin B1. Possibly, palmitic acid induces apoptosis by several mechanisms. In line with this explanation, it has been shown in neonatal rat cardiac myocytes that ceramide accumulation caused by palmitic acid also may occur independent of events that lead to caspase 3–like activation (13,14).

Ceramide induces cytochrome c release from isolated rat liver mitochondria (34). In our study, we demonstrated that palmitic acid induces cytochrome c release also from mitochondria of cardiomyocytes. Thus, our results attest that the mitochondrion is an important target for palmitic acid–induced apoptosis in adult rat cardiac cells.

In neonatal rat cardiomyocytes, high glucose concentrations prevented apoptosis induced by hypoxia (20), although in adult cardiomyocytes, we did not observe any protective effect of glucose against palmitic acid–induced DNA fragmentation. Possibly, neonatal cardiomyocytes respond differently to changes in glucose concentration than postmitotic adult cardiac cells. This hypothesis is supported by the changes in substrate metabolism that occur during the perinatal period, when cardiomyocytes shift from predominant nonoxidative glucose utilization to predominant fatty acid oxidation (35). Alternatively, the protective effect of glucose may be limited to apoptotic pathways other than the one induced by palmitic acid.

Adult cardiomyocytes in long-term culture undergo drastic morphological transitions (36,37). After attachment, the contractile and cytoskeletal structures are almost completely degraded. This is followed by subsequent regeneration of the myofibrillar apparatus. Cells resume spontaneous beating within 1 week in culture. In this context, it is of interest that high glucose concentrations adversely affected the contractile structure not only when added to the cardiomyocytes in the early phase of culture, at a stage of reorganization of myofibrils, but also after 9 days when myofibril formation is mostly completed.

The mechanism that leads to the destruction of the myofibrils caused by elevated glucose concentrations is not clear. Aminoguanidine failed to prevent the deleterious effect of glucose on the myofibrils, suggesting that formation of nitric oxide and/or advanced glycation end products do not play a major role. Blocking de novo synthesis of ceramide from palmitic acid with fumonisin B1 prevented myofibrillar destruction. However, destruction of myofibrils by palmitic acid was followed by DNA fragmentation. Therefore, in the case of fatty acid, the destruction of the myofibrils seems to be an early feature of an apoptotic process. Because glucose did not induce apoptosis, it is unlikely that ceramide formation is involved in the glucose-mediated disruption of the myofibrillar apparatus.

At a physiological concentration of 0.5 mmol/l fatty acids (0.25 mmol/l palmitate and oleate), oleic acid inhibits the deleterious effects of palmitic acid. However, when the concentration of the mixture of fatty acids was elevated to levels more typical of that found in uncontrolled diabetes, the protective effect of oleic acid was insufficient to prevent palmitic acid–induced apoptosis completely. Therefore, it is likely that the toxic effects of the saturated palmitic acid do not occur under physiological conditions. However, in conditions associated with elevated free fatty acid levels or with significantly decreased unsaturated fatty acids, cardiac lipotoxicity may occur.

In conclusion, our results suggest that adult rat cardiac cells are sensitive to increased glucose concentrations and to the saturated palmitic acid. Palmitic acid induces apoptosis via de novo ceramide formation and activation of the mitochondrial apoptotic pathway. Both cell nutrients promote loss of myofibrillar organization and sarcomeric disarray, as observed in cardiomyopathy of failing hearts. Thus, gluco- and lipotoxicity may play a pivotal role in the development of diabetic cardiomyopathy. Identification of these other mechanisms in diabetic cardiomyopathy may lead to novel means of preventing the deleterious effects of hyperglycemia and elevated free fatty acids on the heart of diabetic patients.

FIG. 1.

Disruption of myofibrils and modulation of apoptosis by palmitic acid and glucose in adult rat cardiomyocytes. Confocal micrographs of cardiac cells after 10 days in culture triple labeled with the TUNEL assay (alkaline phosphatase) (A, C, and E) as well as with phalloidin-rhodamine for F-actin (red) and with a monoclonal antibody against myomesin (green-yellow) (B, D, and F). A and B: Control (0.08 mmol/l BSA and 10% FCS). C and D: Cardiomyocytes exposed for 18 h to control media containing 0.5 mmol/l palmitic acid. E and F: Cardiomyocytes exposed for 8 days to media containing 33.3 mmol/l glucose. The arrow marks fragmented nuclei that stained positive for the TUNEL reaction.

FIG. 1.

Disruption of myofibrils and modulation of apoptosis by palmitic acid and glucose in adult rat cardiomyocytes. Confocal micrographs of cardiac cells after 10 days in culture triple labeled with the TUNEL assay (alkaline phosphatase) (A, C, and E) as well as with phalloidin-rhodamine for F-actin (red) and with a monoclonal antibody against myomesin (green-yellow) (B, D, and F). A and B: Control (0.08 mmol/l BSA and 10% FCS). C and D: Cardiomyocytes exposed for 18 h to control media containing 0.5 mmol/l palmitic acid. E and F: Cardiomyocytes exposed for 8 days to media containing 33.3 mmol/l glucose. The arrow marks fragmented nuclei that stained positive for the TUNEL reaction.

FIG. 2.

A: Time course of palmitic acid–induced cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid (Pal) for 1, 3, 6, 10, or 18 h or treated with 33.3 mmol/l glucose from day 9 to day 10 in culture (Gluc 1 d), from day 6 to day 10 in culture (Gluc 4 d), from day 2 to day 10 in culture (Gluc 8 d), from day 9 to day 13 in culture (Gluc 4 d late), or in combination with 0.5 mmol/l palmitic acid for 18 h at day 9 of culture (Pal+Gluc 18 h). Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations (100%; in absolute value: 15% TUNEL-positive cardiomyocytes). The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of at least three independent experiments. Cardiomyocytes were isolated from 32 rats. *P < 0.01 relative to control. B: Dose response of palmitic acid– and oleic acid–induced changes in cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.1, 0.25, or 0.5 mmol/l palmitic (Pal) or oleic (Ole) acid alone or in combination (Ole/Pal) for 18 h. Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations (100%; in absolute value: 12% TUNEL-positive cardiomyocytes). The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of three independent experiments. *P < 0.02 relative to control.

FIG. 2.

A: Time course of palmitic acid–induced cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid (Pal) for 1, 3, 6, 10, or 18 h or treated with 33.3 mmol/l glucose from day 9 to day 10 in culture (Gluc 1 d), from day 6 to day 10 in culture (Gluc 4 d), from day 2 to day 10 in culture (Gluc 8 d), from day 9 to day 13 in culture (Gluc 4 d late), or in combination with 0.5 mmol/l palmitic acid for 18 h at day 9 of culture (Pal+Gluc 18 h). Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations (100%; in absolute value: 15% TUNEL-positive cardiomyocytes). The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of at least three independent experiments. Cardiomyocytes were isolated from 32 rats. *P < 0.01 relative to control. B: Dose response of palmitic acid– and oleic acid–induced changes in cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.1, 0.25, or 0.5 mmol/l palmitic (Pal) or oleic (Ole) acid alone or in combination (Ole/Pal) for 18 h. Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations (100%; in absolute value: 12% TUNEL-positive cardiomyocytes). The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of three independent experiments. *P < 0.02 relative to control.

FIG. 3.

Characterization of the effect of palmitic acid on cardiomyocyte death by double fluorescence staining with Annexin-V-FLUOS (green; A and C) and propidium iodide (red; B and D). Adult rat cardiomyocytes after 9 days in culture were exposed for 18 h to control media (control; A and B) and to 0.5 mmol/l palmitic acid (C and D). The arrow Ap marks an apoptotic cell staining positive only for Annexin-V-FLUOS, and the arrows Ne mark a necrotic cell positive for Annexin-V-FLUOS and with nuclei positive for propidium iodide (fluorescence microscopy, ×400).

FIG. 3.

Characterization of the effect of palmitic acid on cardiomyocyte death by double fluorescence staining with Annexin-V-FLUOS (green; A and C) and propidium iodide (red; B and D). Adult rat cardiomyocytes after 9 days in culture were exposed for 18 h to control media (control; A and B) and to 0.5 mmol/l palmitic acid (C and D). The arrow Ap marks an apoptotic cell staining positive only for Annexin-V-FLUOS, and the arrows Ne mark a necrotic cell positive for Annexin-V-FLUOS and with nuclei positive for propidium iodide (fluorescence microscopy, ×400).

FIG. 4.

Quantitative evaluation of the myofibrillar integrity. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid for 18 h (Pal 18 h), or to 33.3 mmol/l glucose for 18 h (Gluc 18 h), or in combination for 18 h (Pal/Gluc 18 h). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.02 relative to control.

FIG. 4.

Quantitative evaluation of the myofibrillar integrity. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid for 18 h (Pal 18 h), or to 33.3 mmol/l glucose for 18 h (Gluc 18 h), or in combination for 18 h (Pal/Gluc 18 h). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.02 relative to control.

FIG. 5.

Time course of palmitic acid–induced myofibrillar degeneration. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid (Pal) for 1, 3, 6, 10, or 18 h. Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.01 relative to control.

FIG. 5.

Time course of palmitic acid–induced myofibrillar degeneration. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (Control) and to 0.5 mmol/l palmitic acid (Pal) for 1, 3, 6, 10, or 18 h. Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.01 relative to control.

FIG. 6.

Time course of glucose-induced myofibrillar degeneration. Adult rat cardiomyocytes in culture were exposed 10 or 13 days to media containing 5.5 mmol/l glucose (Control and Control 4 d late, respectively) or to 33.3 mmol/l glucose from day 9 to day 10 in culture (Gluc 1 d) or from day 6 to day 10 in culture (Gluc 4 d) or from day 2 to day 10 in culture (Gluc 8 d) or from day 9 to day 13 in culture (Gluc 4 d late). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.05 or **P < 0.01 relative to respective controls.

FIG. 6.

Time course of glucose-induced myofibrillar degeneration. Adult rat cardiomyocytes in culture were exposed 10 or 13 days to media containing 5.5 mmol/l glucose (Control and Control 4 d late, respectively) or to 33.3 mmol/l glucose from day 9 to day 10 in culture (Gluc 1 d) or from day 6 to day 10 in culture (Gluc 4 d) or from day 2 to day 10 in culture (Gluc 8 d) or from day 9 to day 13 in culture (Gluc 4 d late). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.05 or **P < 0.01 relative to respective controls.

FIG. 7.

Lack of protection by aminoguanidine from glucose-induced myofibrillar degeneration. Adult rat cardiomyocytes in culture were exposed 10 or 13 days to media containing 5.5 mmol/l glucose (Control and Control 4 d late, respectively) or to 33.3 mmol/l glucose from day 2 to day 10 in culture (Gluc 8 d) or from day 9 to day 13 in culture (Gluc 4 d late) in the absence or presence of 0.5 mmol/l aminoguanidine (AG). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.05 or **P < 0.01 relative to respective controls.

FIG. 7.

Lack of protection by aminoguanidine from glucose-induced myofibrillar degeneration. Adult rat cardiomyocytes in culture were exposed 10 or 13 days to media containing 5.5 mmol/l glucose (Control and Control 4 d late, respectively) or to 33.3 mmol/l glucose from day 2 to day 10 in culture (Gluc 8 d) or from day 9 to day 13 in culture (Gluc 4 d late) in the absence or presence of 0.5 mmol/l aminoguanidine (AG). Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.05 or **P < 0.01 relative to respective controls.

FIG. 8.

Effect of blockade of ceramide synthesis and of exogenous ceramide in cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 or 33.3 mmol/l glucose alone (control) and to 15 μmol/l C2-dihydroceramide (DH-Ceramide) or 15 μmol/l C2-ceramide (Ceramide) or 0.5 mmol/l palmitic acid (Pal) with or without 15 μmol/l fumonisin B1 (Fumo) or 50 μmol/l sphingomyelinase (Sphyngo), for 18 h. Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations at 5.5 mmol/l glucose alone. The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of at least three independent experiments. **P < 0.01 relative to controls.

FIG. 8.

Effect of blockade of ceramide synthesis and of exogenous ceramide in cardiomyocyte DNA fragmentation. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 or 33.3 mmol/l glucose alone (control) and to 15 μmol/l C2-dihydroceramide (DH-Ceramide) or 15 μmol/l C2-ceramide (Ceramide) or 0.5 mmol/l palmitic acid (Pal) with or without 15 μmol/l fumonisin B1 (Fumo) or 50 μmol/l sphingomyelinase (Sphyngo), for 18 h. Results are mean ± SE of the relative number of TUNEL-positive cardiac cells, normalized to control incubations at 5.5 mmol/l glucose alone. The number of cells scored for DNA fragmentation was 600–1,000 for each treatment condition out of at least three independent experiments. **P < 0.01 relative to controls.

FIG. 9.

Effect of blockade of ceramide synthesis and of exogenous ceramide in myofibrillar degeneration. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 or 33.3 mmol/l glucose alone (Control) and to 15 μmol/l C2-dihydroceramide (DH-Ceramide) or 15 μmol/l C2-ceramide (Ceramide) or 0.5 mmol/l palmitic acid (Pal) with or without 15 μmol/l fumonisin B1 (Fumo) or 50 μmol/l sphingomyelinase (Sphyngo), for 18 h. Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.02 or **P < 0.01 relative to controls.

FIG. 9.

Effect of blockade of ceramide synthesis and of exogenous ceramide in myofibrillar degeneration. Adult rat cardiomyocytes after 9 days in culture were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 or 33.3 mmol/l glucose alone (Control) and to 15 μmol/l C2-dihydroceramide (DH-Ceramide) or 15 μmol/l C2-ceramide (Ceramide) or 0.5 mmol/l palmitic acid (Pal) with or without 15 μmol/l fumonisin B1 (Fumo) or 50 μmol/l sphingomyelinase (Sphyngo), for 18 h. Results are mean ± SE of the score attributed as described in research design and methods. The number of cells evaluated was 600–1,000 for each treatment condition out of at least three independent experiments. *P < 0.02 or **P < 0.01 relative to controls.

FIG. 10.

Cytochrome c release from mitochondria of fatty acid–treated cardiac cells. Left: Immunoblotting of cytochrome c (Cyt c) and of adenine nucleotide translocator (ANT) was performed on mitochondrial fractions of cardiomyocytes. After 9 days in culture, cardiac cells were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (control) and to 0.5 mmol/l palmitic acid (Pal) for 6 or 18 h. Both antibodies were blotted on the same membrane after stripping. A representative experiment of three is shown. Right: Confocal micrographs of a cardiac cell after 9 days in culture exposed for 18 h to media containing 0.5 mmol/l palmitic acid. Triple labeling with the TUNEL assay (alkaline phosphatase) as well as with an antibody against adenine nucleotide translocator (red) and with an antibody against cytochrome c (green). The red arrow marks nuclei that stained positive for the TUNEL reaction. The white arrows mark mitochondria that stained positive for the adenine nucleotide translocator but negative for cytochrome c.

FIG. 10.

Cytochrome c release from mitochondria of fatty acid–treated cardiac cells. Left: Immunoblotting of cytochrome c (Cyt c) and of adenine nucleotide translocator (ANT) was performed on mitochondrial fractions of cardiomyocytes. After 9 days in culture, cardiac cells were exposed to media containing 0.08 mmol/l BSA, 10% FCS, and 5.5 mmol/l glucose alone (control) and to 0.5 mmol/l palmitic acid (Pal) for 6 or 18 h. Both antibodies were blotted on the same membrane after stripping. A representative experiment of three is shown. Right: Confocal micrographs of a cardiac cell after 9 days in culture exposed for 18 h to media containing 0.5 mmol/l palmitic acid. Triple labeling with the TUNEL assay (alkaline phosphatase) as well as with an antibody against adenine nucleotide translocator (red) and with an antibody against cytochrome c (green). The red arrow marks nuclei that stained positive for the TUNEL reaction. The white arrows mark mitochondria that stained positive for the adenine nucleotide translocator but negative for cytochrome c.

This work is supported by the Gustave Prévot Foundation. M.Y.D is supported by the Max Cloetta Foundation.

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Address correspondence and reprint requests to Marc Y. Donath, MD, Division of Endocrinology and Diabetes, Department of Medicine, University Hospital, CH-8091 Zurich, Switzerland. E-mail: marc.donath@dim.usz.ch.

Received for publication 4 October 2000 and accepted in revised form 17 May 2001.

BSA, bovine serum albumin; FCS, fetal calf serum; TUNEL, terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling.