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Division of Pharmaceutical Sciences & Center for Cardiovascular Research and Alternative Medicine, 1000 E. University Avenue, Dept. 3375, University of Wyoming, Laramie, WY 82071. E-mail: firstname.lastname@example.org
Objective: Consumption of high-fat diet and alcohol is associated with obesity, leading to enhanced morbidity and mortality. This study was designed to examine the interaction between high-fat diet and the alcohol metabolizing enzyme alcohol dehydrogenase (ADH) on ethanol-induced cardiac depression.
Research Methods and Procedures: Mechanical and intracellular Ca2+ properties were measured in cardiomyocytes from ADH transgenic and Friend Virus-B type (FVB) mice fed a low- or high-fat diet for 16 weeks. Expression of protein kinase B (Akt) and Foxo3a, two proteins essential for cardiac survival, was evaluated by Western blot. Cardiac damage was determined by carbonyl formation.
Results: High fat but not ADH induced obesity without hyperglycemia or hypertension, prolonged time-to-90% relengthening (TR90), and depressed peak shortening (PS) and maximal velocity of shortening/relengthening (± dL/dt) without affecting intracellular Ca2+ properties. Ethanol suppressed PS and intracellular Ca2+ rise in low-fat-fed FVB mouse cardiomyocytes. ADH but not high-fat diet shifted the threshold of ethanol-induced inhibition of PS and ± dL/dt to lower levels. The amplitude of ethanol-induced cardiac depression was greater in the high-fat but not the ADH group without additive effects. Ethanol down- and up-regulated Akt and Foxo3a expression, respectively, and depressed intracellular Ca2+ rise, the effects of which were exaggerated by ADH, high-fat, or both. High-fat diet, but not ADH, enhanced Foxo3a expression and carbonyl content in non-ethanol-treated mice. Ethanol challenge significantly enhanced protein carbonyl formation, with the response being augmented by ADH, high-fat, or both.
Discussion: Our data suggest that high-fat diet and ADH transgene may exaggerate ethanol-induced cardiac depression and protein damage in response to ethanol.
Obesity, if uncorrected, leads to cardiac hypertrophy and compromised myocardial contractile function and energy metabolism, which are responsible for the high cardiovascular morbidity and mortality in overweight and obese individuals (1, 2, 3, 4). In addition to a wide variety of genetic and lifestyle factors, recent evidence has suggested that consumption of alcoholic beverages may be an independent risk factor for obesity (5). Alcohol consumption directly contributes to heart muscle damage and ventricular dysfunction, namely, alcoholic cardiomyopathy, and increased cardiac mortality in alcoholics (6, 7). Several rationales have been postulated for the pathogenesis of alcoholic cardiomyopathy, including toxicity of alcohol, its metabolite acetaldehyde, and fatty acid ethyl esters, as well as oxidative stress (7, 8, 9). Both clinical and experimental observations have shown a role of the ethanol metabolizing enzyme alcohol dehydrogenase (ADH)1 in the onset and progression of alcoholic cardiomyopathy (10, 11, 12), suggesting a role of ADH in alcohol-induced accumulation of free radicals, oxidative stress, and heart disease. Through elevated expression of ADH, the enzyme that hydrolyzes ethanol into acetaldehyde, progression of alcoholic cardiomyopathy may become significantly “accelerated” in both functional and morphological aspects (10, 12, 13, 14). Nevertheless, the impact of obesity on ADH enzyme-facilitated myopathic alteration in response to alcohol intake has not been elucidated. Given the close epidemiological association between body mass and alcohol intake (5, 15), this study was designed to examine the impact of high-fat diet-induced obesity on basal cardiac function and the ADH-induced sensitization of ethanol-elicited depression of cardiac function and the mechanism of action involved, with an emphasis on the cardiac survival factor protein kinase B (Akt) and the pro-apoptotic forkhead transcriptional factor Foxo3a. Levels of the serine-threonine protein kinase Akt and the pro-apoptotic Foxo3a have been shown to be rather sensitive to changes in tissue insulin sensitivity, such as in obesity and diabetes (16, 17). Ethanol has been demonstrated to directly regulate cardiomyocyte survival through phosphorylation of Akt (18), although little information is available for the involvement of Foxo3a. Given the important role of Akt and Foxo3a in cardiomyocyte survival and contractile function (19, 20, 21), these proteins were evaluated in an effort to explore the potential mechanism of action behind the interaction of high-fat diet and ADH, if any, on cardiomyocyte mechanical function. The fat-enriched diet has been commonly used to foster diet-induced prediabetic normotensive or mildly hypertensive obesity (17).
Research Methods and Procedures
ADH Transgenic Mice and High-Fat Diet Feeding
The experimental procedure described here was approved by the University of Wyoming Institutional Animal Use and Care Committee and was in accordance with NIH standards. Production of the ADH transgenic mice was described previously (14). In brief, using albino Friend Virus-B type (FVB) mice, the cDNA for murine Class I ADH was inserted behind the mouse α-myosin heavy chain promoter to achieve cardiac-specific overexpression. This cDNA was selected because Class I ADH is most efficient in the oxidation of ethanol. A second transgene with a cDNA encoding tyrosinase was co-injected with ADH to produce coat color pigmentation (dark gray) in albino mice. The ADH-positive mice displayed an ∼40-fold increase in cardiac ADH protein expression associated with a 3- to 5-fold increase in ADH enzymatic activity (which converts ethanol into acetaldehyde) (14, 22). Both ADH transgenic and FVB wild-type male mice (3 months old) were randomly assigned to either low-fat (10% of total calorie) or high-fat (45% of total calorie) diets (Research Diets, Inc., New Brunswick, NJ) for 16 weeks. The high-fat diet was calorically rich (4.83 kcal/g vs. 3.91 kcal/g in low-fat diet) due to higher fat composition. However, the two diets possessed similar nutrient composition (17). Mice were housed in individual cages in a climate-controlled environment (22.8 ± 2.0 °C, 45% to 50% humidity), with a 12/12-light/dark cycle, and free access to diets as well as water. At the end of 16 weeks of dietary feeding, fasting (12 hours) blood glucose and systolic blood pressure were measured by an Accu-Chek III glucose analyzer (F. Hoffmann-La Roche Ltd, Basel, Switzerland), and a semi-automated, amplified tail cuff device (IITC, Inc., Woodland Hills, CA), respectively. For systolic blood pressure measurement, mice were conditioned to the restraining cylinders, pre-warmed at 29 °C for 8 to 10 minutes to facilitate the tail blood flow before blood pressure was measured. The mean of three readings was used as the systolic blood pressure value for each mouse. For the acute ethanol challenge experiment used for biochemical and Western analyses, mice, following 16 weeks of low- or high-fat dietary intake, were injected intraperitoneally with ethanol (3 g/kg per day) consecutively for 3 days and were killed 24 hours after the last ethanol injection for cardiac tissue collection.
Murine Cardiomyocyte Isolation Procedures
After ketamine/xylazine (3:1, 1.32 mg/kg body weight) sedation, mouse hearts were removed and perfused with Krebs-Henseleit bicarbonate buffer containing (in mM) 118 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 25 NaHCO3, 10 HEPES, and 11.1 glucose, with 5% CO2-95% O2. Hearts were subsequently digested with a Krebs-Henseleit bicarbonate buffer containing 223 U/mL collagenase D (Boehringer Mannheim, Indianapolis, IN) for 20 minutes. After perfusion, left ventricles were removed and minced before being filtered. Extracellular Ca2+ was slowly added back to 1.25 mM. Myocytes with obvious sarcolemmal blebs or spontaneous contractions were not used (22). Myocytes were used within 6 hours of isolation.
Mechanical properties of cardiomyocytes were assessed using a SoftEdge MyoCam system (IonOptix, Milton, MA) as described (22). In brief, cells were placed in a chamber mounted on the stage of an inverted microscope (Olympus IX-70; Olympus America, Inc., Center Valley, PA) and superfused (∼2 mL/min at 25 °C) with a buffer containing (in mM) 131 NaCl, 4 KCl, 1 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES at pH 7.4. The cells were field stimulated to contract at a frequency of 0.5 Hz. Changes in cell length during shortening and relengthening were captured and converted to digital signal. The myocyte being studied was rapidly scanned with a camera at 120 Hz to ensure recording with good fidelity. Cell shortening and relengthening were assessed using the following indices: peak shortening (PS), time-to-PS (TPS), time-to-90% relengthening (TR90), and maximal velocities of shortening and relengthening (± dL/dt). In the case of altering stimulus frequency from 0.1 Hz to 5.0 Hz, the steady state contraction of myocyte was achieved (usually after the first 5 to 6 beats) before PS was recorded.
Intracellular Ca2+ Fluorescence Measurement
A cohort of cardiomyocytes was loaded with fura-2/AM (0.5 µM) for 10 minutes at 25 °C, and fluorescence measurements were recorded with a dual-excitation fluorescence photomultiplier system (IonOptix) as previously described (22). Myocytes were imaged through an Olympus IX-70 Fluor oil objective. Cells were exposed to light emitted by a 75W lamp and passed through either a 360 or a 380 nm filter (bandwidths were ±15 nm) while being stimulated to contract at 0.5 Hz. Fluorescence emissions were detected between 480 and 520 nm by a photomultiplier tube after illuminating the cells first at 360 nm for 0.5 seconds and then at 380 nm for the duration of the recording protocol (333-Hz sampling rate). The 360-excitation scan was repeated at the end of the protocol, and qualitative changes in intracellular Ca2+ concentration were inferred from the ratio of the fura-2 fluorescence intensity at the 2 wavelengths. Fluorescence decay time (both single and bi-exponential decay rates) was also measured as an indication of intracellular Ca2+ clearing rate.
Western Blot Analysis of Foxo3a and Akt
The total protein was prepared as described previously (22). In brief, left ventricles were rapidly removed and homogenized in a lysis buffer containing 20 mM Tris (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton, 0.1% sodium dodecyl sulfate, and 1% protease inhibitor cocktail. Samples were then sonicated for 15 seconds and centrifuged at 12,000g for 20 minutes at 4 °C. The protein concentration of the supernatant was evaluated using the Protein Assay Reagent (Bio-Rad Laboratories, Inc., Hercules, CA). Equal amounts (50 µg protein/lane) of protein and pre-stained molecular weight marker (Gibco-BRL, Gaithersburg, MD) were loaded onto 15% sodium dodecyl sulfate-polyacrylamide gels in a mini-gel apparatus (Mini-PROTEAN II, Bio-Rad), separated, and transferred to nitrocellulose membranes (0.2 µm pore size, Bio-Rad Laboratories, Inc.). Membranes were incubated for 1 hour in a blocking solution containing 5% non-fat milk in Tris-buffered saline before being washed in Tris-buffered saline and incubated overnight at 4 °C with anti-Akt (1:1000), Foxo3a (1:1000), and anti-β-actin (1:5000, used as loading control) antibodies. Anti-Akt and anti-β-actin antibodies were purchased from Cell Signaling Technology (Beverly, MA), and anti-Foxo3a antibody was obtained from Upstate (Lake Placid, NY). After incubation with the primary antibodies, blots were incubated with either anti-mouse or anti-rabbit immunoglobulin G horseradish peroxidase-linked antibodies at a dilution of 1:5000 for 60 minutes at room temperature. Immunoreactive bands were detected using the Super Signal West Dura Extended Duration Substrate (Pierce, Milwaukee, WI). The intensity of bands was measured with a scanning densitometer (model GS-800; Bio-Rad) coupled with Bio-Rad PC analysis software. For each protein of interest, the Western blotting was repeated at least twice to confirm the accuracy of the data. Only one myocardial sample was used from each mouse for Western blot analysis. All optical density readings for a given sample were normalized to that of the β-actin.
Protein Carbonyl Assay
To assess oxidative protein damage, the carbonyl content of protein extracted from hearts was determined as described (12). Briefly, proteins were extracted and minced to prevent proteolytic degradation. Nucleic acids were eliminated by treating the samples with 1% streptomycin sulfate for 15 minutes, followed by a 10-minute centrifugation (11,000g). Protein was precipitated by adding an equal volume of 20% trichloroacetic acid to protein (0.5 mg) and centrifuged for 1 minute. The trichloroacetic acid solution was removed and the sample resuspended in 10 mM 2,4-dinitrophenylhydrazine solution. Samples were incubated at room temperature for 15 to 30 minutes. After the addition of 500 µL of 20% trichloroacetic acid, samples were centrifuged for 3 minutes. The supernatant was discarded, and the pellet was washed in ethanol:ethyl acetate and allowed to incubate at room temperature for 10 minutes. The samples were centrifuged again for 3 minutes, and the ethanol:ethyl acetate steps were repeated 2 more times. The precipitate was resuspended in 6-mol/L guanidine solution and centrifuged for 3 minutes and the insoluble debris removed. The maximum absorbance (360 to 390 nm) of the supernatant was read against appropriate blanks (water, 2 mol/L HCl), and the carbonyl content was calculated using the molar absorption coefficient of 22,000 M−1cm−1.
For in vitro ethanol exposure, myocytes (fura-2 loaded or non-loaded) from mice without the intraperitoneal ethanol challenge were first allowed to contract at a stimulation frequency of 0.5 Hz for 5 minutes to ensure steady state (myocytes with rundown >10% were not studied further) before perfusing with ethanol (80 to 640 mg/dL) containing contractile buffer. A 3-minute interval was allowed between the ethanol doses. Our previous experience indicated that ethanol-induced inhibition of cell shortening remains stable for up to 30 minutes (10, 23).
Data are means ± standard error. Statistical comparisons were performed by ANOVA followed by Newman-Keuls post hoc test. Significance was defined as p < 0.05.
General Feature of FVB and ADH Mice Fed With Low- and High-Fat Diets
Although the high-fat diet was calorically rich (4.83 kcal/g vs. 3.91 kcal/g in low-fat diet) due to higher fat composition, the average daily caloric intake was comparable between the low- and high-fat diet-fed FVB and ADH groups (low-fat FVB: 8.99 kcal/mouse per day; high-fat FVB: 9.42 kcal/mouse per day; low-fat ADH: 8.60 kcal/mouse per day; and high-fat ADH: 9.30 kcal/mouse per day). This similarity in total caloric intake between the low- and high-fat diet groups is consistent with the previous report from our laboratory (17). High-fat feeding significantly increased body and organ weights compared with low-fat-fed mice in both the FVB and ADH strain. The size of the hearts, livers, and kidneys (organ weight normalized to body weight), however, was unaffected by high-fat diet feeding. ADH transgene did not affect body or organ weights, regardless of the diet regimen. Levels of fasting blood glucose and systolic blood pressure were unaffected by either high-fat diet feeding or the ADH transgene (Table 1).
Table 1. . General features of FVB and ADH mice fed an LF or HF diet for 16 weeks
FVB-LF (n = 18)
FVB-HF (n = 15)
ADH-LF (n = 16)
ADH-HF (n = 24)
FVB, Friend Virus-B type; ADH, alcohol dehydrogenase; LF, low-fat; HF, high-fat. Data are means ± standard error; n = number of animals.
Acute Effects of Ethanol on Cell Shortening and Intracellular Ca2+ Transients
To determine if high-fat diet or ADH transgene affects ethanol-induced depression of cardiomyocyte shortening, concentration-dependent responses were constructed for ethanol (0 to 640 mg/dL) in myocytes from FVB and ADH mice fed with low- or high-fat diet. Either ADH transgene or high-fat diet alone did not affect the resting cell length. However, the joint effect of ADH transgene and high-fat diet significantly increased the resting cell length (Figure 1A). High-fat diet reduced PS and maximal velocity of shortening and relengthening (± dL/dt) and prolonged TR90 without affecting TPS in FVB but not ADH mice. Similar to results in previous reports (10, 13), ADH transgene itself did not affect the mechanical indices tested. Acute ethanol exposure did not alter resting cell length but significantly depressed PS and ± dL/dt in all four mouse groups. Consistent with our previous reports (10, 13), ADH transgene shifted the threshold of ethanol-induced inhibition of PS and ± dL/dt to the left. The threshold of ethanol-induced inhibition of PS and ± dL/dt was shifted from 80 to 120 mg/dL in the FVB group to <80 mg/dL in the ADH group. Interestingly, ethanol-induced prolongation of TPS and TR90 (at 240 and 640 mg/dL) in FVB myocytes was not seen in ADH myocytes. High-fat diet feeding did not significantly affect the pattern of ethanol-induced inhibition of PS and ± dL/dt. However, the maximal inhibition of PS and ± dL/dt in response to ethanol was greater in high-fat-fed mice than in the low-fat group. The threshold of ethanol-elicited inhibition of PS and ± dL/dt was comparable (80 to 120 mg/dL) between the low- and high-fat diet FVB groups. Interestingly, cardiomyocytes from high-fat-fed mice were less sensitive to an ethanol-induced effect of TPS and TR90, possibly due to an elongated TR90 or a non-significantly prolonged TPS. Furthermore, the combination of ADH transgene and high-fat diet did not produce any additional effect on myocyte mechanical indices compared with high-fat diet alone. However, the joint depressant effect of high-fat diet and ADH transgene on PS and ± dL/dt was significantly greater that that elicited by ADH transgene alone (Figure 1). Consistent with its response on cell shortening, ethanol significantly depressed the electrically stimulated rise in intracellular Ca2+ fluorescence in a concentration-dependent manner (120 and 640 mg/dL) in FVB myocytes, with a threshold higher than 120 mg/dL. Both high-fat diet and ADH transgene exaggerated ethanol-induced inhibitory response on intracellular Ca2+ rise with a threshold lower than 120 mg/dL. The combination of high-fat diet and ADH transgene did not produce any further inhibition in response to ethanol compared with either maneuver alone. Neither high-fat diet nor ADH transgene itself altered the resting intracellular Ca2+ levels and intracellular Ca2+ decay rate (mono- or bi-exponential). Nonetheless, the combination of high-fat diet and ADH transgene elevated resting intracellular Ca2+ without affecting intracellular Ca2+ decay rate (mono- or bi-exponential). Ethanol exposure slowed the bi-exponential intracellular Ca2+ decay rate at 640 mg/dL but failed to affect resting intracellular Ca2+ levels or mono-exponential intracellular Ca2+ decay rate in low fat-fed FVB myocytes. The high-fat diet did not affect the pattern of ethanol-induced responses. To the contrary, ADH transgene unmasked ethanol-induced decrease in resting intracellular Ca2+ and prolongation of intracellular Ca2+ decay rate (mono- or bi-exponential). Surprisingly, the combination of high-fat diet and ADH transgene abrogated ethanol-induced decreases in intracellular Ca2+ decay rate (mono- or bi-exponential) by either maneuver alone without affecting the ADH-induced depression of intracellular Ca2+ in response to 640 mg/dL ethanol (Figure 2). Rodent hearts contract at very high frequencies, whereas our mechanical and intracellular Ca2+ evaluations were conducted at 0.5 Hz. To evaluate the impact of high-fat diet and ADH transgene on cardiomyocyte contractile function under physiological frequencies, we increased the stimulating frequency up to 5.0 Hz (300 beats/min) and recorded steady-state peak shortening. Cells were initially stimulated to contract at 0.5 Hz for 5 minutes to ensure steady state before increasing the frequency response from 0.1 to 5.0 Hz. PS obtained at all recording frequencies was normalized to that at 0.1 Hz of the same myocyte. Figure 3 shows comparable negative staircases in PS with increasing stimulus frequency between the FVB and ADH mice fed a low-fat diet. However, myocytes from high-fat diet-fed FVB and ADH mice displayed a significantly enhanced depression in PS at all frequencies tested, suggesting possibly reduced intracellular Ca2+ cycling after high-fat diet feeding.
Akt, Foxo3a Protein Expression, and Protein Carbonyl Formation After Ethanol Exposure
To examine possible signaling mechanism(s) involved in ethanol-induced cardiac contractile depression in low- and high-fat-fed FVB and ADH mice, all 4 groups of mice were injected intraperitoneally with ethanol (3 g/kg per day) for 3 consecutive days before cardiac tissues were collected for gel electrophoresis and protein carbonyl determination. The data in Figure 4 reveal that expressions of the cardiac survival factor Akt and the pro-apoptotic factor Foxo3a were significantly down- and up-regulated, respectively, by ethanol treatment among all groups tested. Either high-fat diet or ADH transgene significantly exaggerated ethanol-induced down-regulation of Akt and up-regulation of Foxo3a without any additive effect between the two. The levels of Akt were similar among non-ethanol-treated FVB and ADH mice fed with either low- or high-fat diet. However, consistent with our earlier report (17), high-fat diet, but not ADH transgene, up-regulated expression of Foxo3a in non-ethanol-treated mice. Analysis of the protein carbonyl formation, an index of protein damage, depicted findings similar to those for Foxo3a. While high-fat diet, but not ADH transgene, significantly up-regulated protein carbonyl formation in non-ethanol-treated mice, ethanol challenge significantly enhanced cardiac protein carbonyl formation in all four groups tested. Either high-fat diet or ADH transgene significantly exaggerated ethanol-induced protein damage without any additive effect between the two.
The major findings of this study were that high-fat diet feeding but not the ADH transgene induced obesity, prolonged relaxation duration (TR90), and depressed cardiac contractility and maximal rate of force development/decline (± dL/dt) without affecting the systolic duration (TPS) and intracellular Ca2+ properties. Consistent with previous findings (10, 13, 22), ethanol suppressed PS and intracellular Ca2+ rise in low-fat-fed FVB mouse cardiomyocytes. ADH transgene shifted the threshold of ethanol-induced inhibition of PS and ± dL/dt to the left, depicting a “sensitizing effect” of the ADH transgene to ethanol exposure. Although a high-fat diet failed to shift the threshold of effectiveness for ethanol-induced cardiac depression, the degree of ethanol-induced cardiac depression of PS and ± dL/dt was greater in high-fat-fed FVB mice and ADH mice. There was no additive effect between high-fat diet feeding and ADH transgene. Furthermore, ethanol dampened the rise in intracellular Ca2+ in response to electrical stimuli, the effect of which was exacerbated by high-fat diet, ADH transgene, or both. Data from the immunoblots revealed that intraperitoneal administration of ethanol significantly down- and up-regulated Akt and Foxo3a expression, respectively. Carbonyl formation was also significantly elevated in ethanol-treated mice. Similar to intracellular Ca2+ responses, ADH, high-fat feeding, or both significantly augmented ethanol-induced effects on Akt, Foxo3a, and carbonyl formation. These results suggest that high-fat diet-induced obesity might augment the ethanol-induced cardiac depression and alteration of key anti-/pro-apoptotic proteins, in a manner similar to its metabolizing enzyme ADH. Data presented in this work further indicated that high-fat diet did not significantly interact with the ADH enzyme on ethanol-induced cardiac mechanical and intracellular Ca2+ changes, as well as Akt, Foxo3a, and cardiac protein damage. These findings seem to favor the notion that the two maneuvers may share common pathway(s) leading to cell/tissue injury and cardiac depression.
High-fat diet intake is commonly linked to dyslipidemia, insulin resistance, obesity, and type 2 diabetes (24, 25), all of which trigger cardiac contractile dysfunction via sympathetic activation and renin-angiotensin system stimulation (1, 4). Results from the current study revealed compromised cardiomyocyte contractile function (depressed PS and ± dL/dt and prolonged TR90) and reduced stress tolerance at higher stimulating frequencies after high-fat diet intake, consistent with previous findings from both human and experimental obesity (4, 17, 24, 25, 26, 27). The 16-week high-fat diet feeding did not significantly affect systolic blood pressure and fasting blood glucose, ruling out the contribution of concomitant hypertension and diabetes mellitus. The relatively normal or less affected intracellular Ca2+ homeostasis in conjunction with impaired myocyte mechanical properties in high-fat diet-fed mouse myocytes seems to indicate the presence of reduced myofilament Ca2+ sensitivity after chronic high-fat diet intake. This is supported by the observation of exaggerated PS decline at higher stimulus frequencies in high-fat-fed mouse myocytes, indicating dampened intracellular Ca2+ cycling capacity. The fact that ADH transgene masked high-fat diet-induced mechanical dysfunction without eliciting any effect in low- or high-fat-fed mice is puzzling. Although data from our laboratory did not favor any innate cardiac contractile effect from cardiac specific overexpression of ADH (10, 13), it is possible that the transgene may interact with certain cardiac contractile or signaling pathways, which may be turned on by high-fat diet feeding, although further study is warranted. This seems to be consistent with our recent observation that ADH transgene failed to alter cardiomyocyte contractile and intracellular Ca2+ homeostasis in young mice but was able to improve the aging-associated diastolic defect (28).
Impaired cardiac contractile function and interrupted intracellular Ca2+ handling are perhaps the most significant manifestations of ethanol exposure-induced cardiac depression (29, 30, 31). Data from the current study revealed that cardiomyocyte contractile and intracellular Ca2+ properties were drastically depressed by ethanol exposure in all mouse groups tested. In the current study, ethanol prolonged the duration of both shortening and relengthening (TPS and TR90) in conjunction with depressed PS and ± dL/dt in low-fat-fed FVB mouse myocytes, consistent with our earlier study (10). ADH transgene sensitized cardiomyocytes from both low- and high-fat diet groups to ethanol exposure with a leftward shift in the threshold of ethanol effectiveness. Although high-fat diet failed to alter the threshold of ethanol-induced cardiac effects, the degree of ethanol-induced cardiac depression on PS and ± dL/dt was much greater in high-fat-fed FVB mice and ADH mice. High-fat diet and ADH transgene exacerbated the ethanol-induced inhibition of rise in intracellular Ca2+ in response to electrical stimuli. Western blot analysis revealed that intraperitoneal administration of ethanol significantly down- and up-regulated Akt and Foxo3a expression, respectively. Carbonyl formation was also significantly elevated in ethanol-treated mice. These data suggest that ethanol-elicited decrease in the anti-apoptotic protein Akt and rise in pro-apoptotic transcription factor Foxo3a may contribute to ethanol-induced cardiac dysfunction and protein damage (evaluated by carbonyl formation). It is worth mentioning that there was little additive effect between high-fat diet and ADH on cell shortening, intracellular Ca2+ homeostasis, Akt, Foxo3a, and carbonyl formation, suggesting that high-fat diet and ADH transgene may exacerbate the ethanol-induced cardiac responses through a somewhat common pathway, such as apoptotic cascades. Interestingly, data from this study also revealed that high-fat diet only, but not ADH, enhanced Foxo3a expression and carbonyl formation in the absence of ethanol treatment, suggesting the presence of apparent discrepancies in cell injury between the two maneuvers. Further study is warranted to elucidate the stress signaling mechanisms involved in high-fat diet and ADH enzyme-induced exaggeration of the ethanol toxicity.
ADH belongs to a group of dimeric zinc-containing enzymes with a subunit molecular weight of 40 kDa. Six classes (Class I to VI) of ADH have been identified (32). Class I ADH is a sub-family in which most polymorphic alleles may be found in different ethnic populations, including whites, blacks, and Asians (32). Most of the current data on these ADH gene loci are somewhat restricted to the areas of metabolism, toxicity, and alcohol addiction, with relatively limited knowledge on cardiovascular health (32, 33). It has been shown that expression or activity of ADH may be up-regulated by hypoxia and the ethylene-releasing compounds, such as grape berries and 2-chloroethylphosphonic acid (34, 35), which may cause certain clinical concerns given the correlation between ADH expression and risk of alcohol complications or Alzheimer's disease (10, 12, 36).
In summary, this study suggests that high-fat diet-induced obesity may augment the ethanol-induced cardiac depression and alteration of key anti-/pro-apoptotic proteins, in a manner similar to its metabolizing enzyme, ADH. It is possible that high-fat diet intake and ADH transgene may share somewhat similar pathway(s) in exacerbation of the ethanol-induced cell injury and cardiac depression. Future in-depth study of the impact of high-fat diet intake and obesity on alcoholic cardiomyopathy and other alcoholic complications should shed some light on the epidemiological link between obesity and chronic alcoholism and associated organ complications.
The author thanks Feng Dong from University of Wyoming for technical assistance. The ADH founder mice were kindly provided by Paul N. Epstein from University of Louisville, Louisville, KY. This work was supported in part by NIH Grants R15 AA13575 and R01 AA13412.
Nonstandard abbreviations: ADH, alcohol dehydrogenase; Akt, protein kinase B; FVB, Friend Virus-B type; PS, peak shortening; TPS, time-to-PS; TR90, time-to-90% relengthening; ± dL/dt, maximal velocities of shortening and relengthening.