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Am J Physiol Heart Circ Physiol 291: H1047-H1057, 2006. First published April 28, 2006; doi:10.1152/ajpheart.00214.2006
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Biphasic changes in cardiac excitation-contraction coupling early in chronic alcohol exposure

Gary L. Aistrup,1 James E. Kelly,2 Mariann R. Piano,3 and J. Andrew Wasserstrom2

1Departments of Molecular Pharmacology and Biological Chemistry and 2Medicine, Division of Cardiology, Northwestern University, Feinberg School of Medicine; and 3Department of Medical-Surgical Nursing, University of Illinois at Chicago College of Nursing, Chicago, Illinois

Submitted 27 February 2006 ; accepted in final form 26 April 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Although the negative inotropic effects of both acute and chronic ethanol (EtOH) exposure are well known, little is known concerning the acute-to-chronic transition of such effects. In this study, our objective was to address this question by detailing the effects that acute EtOH exposure induces on cellular excitation-contraction (EC) coupling and, subsequently, comparing whether and how such changes translate to the early chronic EtOH condition in a rat model known to develop alcohol-induced cardiomyopathy. Acute EtOH exposure, as formerly reported, indeed induced dose-dependent negative inotropic changes in cellular EC coupling, manifest as reductions in cell shortening, Ca2+ transient amplitude, Ca2+ decay rate, and sarcoplasmic reticulum Ca2+ content of isolated rat ventricular cardiac myocytes. Supplementary to this, we found Ca2+ spark character not to be significantly affected by acute EtOH exposure. In contrast, the results obtained from cardiac myocytes isolated from rats fed a diet containing ~9% (vol/vol) EtOH for 1 mo revealed changes in these parameters reflecting positive inotropy, whereas at 3 mo, these parameters again reflected negative inotropy similar but not identical to that induced by acute EtOH exposure. No significant changes were evident at either 1- or 3-mo chronic EtOH administration in echocardiographic parameters known to be perturbed in alcoholic cardiomyopathy (ACM), thus indicating that we were examining an asymptomatic stage in chronic EtOH administration consistent with an acute-to-chronic transition phase. Continued study of such transition-phase events should provide important insight into which molecular-cellular components of EC coupling play pivotal roles in EtOH-induced disease processes, such as ACM.

rat cardiac myocytes; calcium transients; calcium sparks; confocal calcium imaging

IT HAS LONG BEEN ESTABLISHED that ethanol (EtOH) impairs cardiac contractility, which, as demonstrated by numerous studies, occurs as a result of an impairment of excitation-contraction (EC) coupling (51). Indeed, excessive long-term EtOH consumption produces left ventricular dysfunction and can lead to a dilated cardiomyopathy referred to as alcoholic cardiomyopathy (ACM) (12, 25, 42). A large proportion of alcohol-dependent individuals develops systolic dysfunction and ACM. However, the point at which these abnormalities appear during the course of an individual's lifetime of drinking, such that the abnormalities can be called ACM, is not well established and is highly individualized (4, 26, 31, 48, 50, 53). In fact, not all alcoholics develop symptomatic ACM (13). Although much is known about the acute and late-stage chronic effects of EtOH, little has been delineated regarding the changes in cardiac myocyte function during the transition period where acute EtOH exposure becomes short-term chronic EtOH exposure, particularly at the cellular level. This acute-to-chronic transition period likely encompasses EC coupling events key to understanding the basis for the development and onset variability of ACM and other maladies induced by chronic EtOH consumption.

The objective of the present study was to investigate the events underlying the acute-to-chronic EtOH transition. To this end, we examined and compared the acute and short-term (1 and 3 mo, respectively) effects of EtOH exposure on cellular EC coupling in isolated rat ventricular cardiac myocytes, i.e., cell shortening and Ca2+ transients, and, in addition, we assessed for the first time the effects of acute EtOH exposure at the subcellular level of individual sarcoplasmic reticulum (SR) Ca2+ release units detected as small spontaneous burst of localized fluorescence (Ca2+ sparks). Echocardiographic parameters at 1- and 3-mo EtOH exposure were also measured to identify potential in vivo correlates to our cellular in vitro findings.

Our results indicated that, as expected, acute EtOH exposure induced negative inotropic effects in isolated rat ventricular cardiac myocyte EC coupling manifested as reduced cell shortening and diminished Ca2+ transients. Although, somewhat unexpectedly, we found no significant effect of acute EtOH on spark characteristics. Our main and unique finding, however, was that the negative inotropic effects apparent in cellular EC coupling upon acute EtOH exposure were not apparent at 1-mo chronic EtOH exposure, the latter of which were actually indicative of positive inotropy. However, at 3-mo chronic EtOH exposure, there was a recurrence of negative inotropic effects manifest in cellular EC coupling resembling but not equivalent to acute exposures of EtOH at high concentrations.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents and Solutions

All chemicals used to make the physiological salt solutions were purchased from Sigma-Aldrich (St. Louis, MO) and were of analytical grade or better. Absolute (200 proof) nondenatured EtOH was purchased from Pharmco Products (Brookfield, CT) and maintained in glass bottles to avoid otherwise possible contamination by leachable monomers, plasticizers, and/or stabilizers from plastic containment.

Experimental Groups and Ethanol Consumption Protocol

The animal usage procedures described in this and the following three sections all conformed to National Institutes of Health guidelines and were approved by Northwestern University and the University of Illinois at Chicago Animal Care and use Committees.

Male Sprague-Dawley rats (beginning weight, 248–263 g) were used in all studies. After arrival, rats were housed (3 rats/cage) in a temperature- (21–23°C) and light-controlled (0600–1800 h) environment for a period of 6 days. After this equilibration period, animals were divided into the following groups. One group of rats (n = 5) was used for acute EtOH studies and therefore received a standard chow and water diet. For sustained in vivo EtOH exposure, animals were randomly assigned to control-fed (CNTRL, n = 8) or EtOH-fed (ETOH, n = 8) groups, which were maintained on the CNTRL or ETOH protocol for 1 and 3 mo. Both CNTRL and ETOH groups received the nutritionally complete Lieber-DeCarli 1982 liquid diet formulation (28). In the EtOH diet, the animals received 9% (vol/vol) EtOH, whereas in the control diet, maltose-dextrin was substituted isocalorically for EtOH (32, 43). The CNTRL group was pair-fed with the ETOH group. Dietary consumption was monitored daily, and fresh diet was provided each day between 1600 and 1800 h.

Determinations of Blood Alcohol Concentration

Blood alcohol concentrations (BACs) of chronic ETOH rats were measured by collecting ~1 ml of blood samples from each ETOH rat via tail-vein bleed [rats lightly anesthetized with ketamine (50 mg/kg ip) and xylazine (5 mg/kg ip)] every 2 wk and at the time of death in CNTRL and ETOH animals. BACs of each sample were subsequently determined by using an ADH/NAD UV-spec assay (Ethanol-L3K; Diagnostic Chemicals, Oxford, CT). At death, the mean BACs were 0.15 ± 0.03% (wt/vol) EtOH for the 1-mo EtOH-fed (1-mo-ETOH) group and 0.19 ± 0.04% (wt/vol) EtOH for the 3-mo-EtOH-fed (3-mo ETOH) group, which are similar to those often maintained by humans suffering from alcoholism. (1, 2, 29).

Echocardiography

To determine whether cardiac function was altered during chronic EtOH consumption, echocardiograms were performed after 1 and 3 mo in CNTRL and ETOH rats. All echocardiograms were performed by the same sonographer using the Sequoia C256 Echocardiography System (Acuson, Mountain View, CA) and a 14L/8 frequency transducer (15.0-MHz acquisition). Before the procedure, animals were anesthetized with ketamine (40–50 mg/kg ip) and xylazine (5 mg/kg ip). The transducer was placed on the left thorax, and M-mode recordings were made by directing the ultrasound beam at the midpapillary muscle level. The measurements listed below were obtained after a well-defined, continuous interface of the anterior and posterior walls were visualized.

According to the methods of the American Society of Echocardiography and other investigations in animal models, the left ventricular internal dimensions at end diastole and end systole (LVIDd and LVIDs, respectively), interventricular septum at end diastole (IVSd), and left ventricular posterior wall at end diastole (LVPWd) were obtained via the leading-edge method (46, 47). All parameters were measured with electronic calipers, and mean calculations were obtained from three or more consecutive cardiac cycles. Intraobserver variability for all echocardiographic parameters ranged from 0 to 10% (mean, 7%). The following parameters were calculated from the obtained parameters according to accepted formulas (37):

Formula 1(1)

Formula 2(2)

Formula 3(3)

Cardiac Myocyte Isolation

Ventricular cardiac myocytes were isolated from rats using procedures modified from, but based on, those previously described (19, 27, 34). Briefly, hearts were excised from heparinized (2,500 U/kg ip) rats under halothane (to effect) or ketamine + xylazine (87 mg/kg + 13 mg/kg ip) anesthesia, mounted on a Langendorff perfusion apparatus, and perfused with nominally Ca2+-free HEPES-buffered Tyrode solution for 5 min at 37°C. The heart was subsequently perfused with the 40 µmol/l Ca2+-containing Tyrode solution + 1 mg/ml collagenase (Worthington type II or Boehringer Mannheim type B) + 1 mg/ml BSA until it became flaccid (10–15 min). During the final 4 min of this latter perfusion, small aliquots of stock CaCl2 solution were added incrementally until the final Ca2+ concentration was 1 mmol/l. After this perfusion, the left ventricle was dissected from the rest of the heart, minced, gently shaken, triturated in 1 mmol/l Ca2+-containing Tyrode + 1 mg/ml collagenase + 2 mg/ml BSA, and filtered over a 300-µm nylon mesh to separate the cells from the digested collagen matrix. The resultant cell suspension was rinsed several times with progressive increases in [Ca2+] to 1.8 mmol/l and, finally, resuspended in medium 199 (GIBCO), aliquoted into 15-ml Petri dishes, and maintained in a humidified 5% CO2-95% air cell-culture incubator at 37°C until use (typically within 12 h).

Cardiac myocyte isolations from chronic ETOH rats were performed identically, except that all isolation solutions were made to contain a concentration of EtOH equivalent to the average BAC that the rat had maintained 2 wk previously at the time of death so as to provide an equivalent in vitro EtOH-containing environment and to avoid complications associated with acute alcohol withdrawal. After isolation, the cells isolated from ETOH rats were maintained in Tyrode or medium 199 also containing the same concentration of EtOH.

Image Acquisition of Cell Shortening, Ca2+ Transients, and Sparks

All experiments were conducted at room temperature (21–23°C). Cells were loaded via incubation in HEPES-buffered Tyrode solution (±EtOH, see below) containing 5 µmol/l of the acetoxymethyl ester of the fluorescent Ca2+ indicator fluo-4 (Molecular Probes) for 20–30 min. The cells were then washed with normal Tyrode solution, and aliquots were gently pipetted into a perfusion chamber mounted on the stage of an inverted laser-scanning confocal microscope (Axiovert 100M coupled to an LSM 510 system utilizing a 25-mW argon laser, Carl Zeiss) where they settled on the lamanin-coated glass bottom of the chamber. The cells were then constantly superfused with normal Tyrode solution at room temperature (21–23°C) at a 5 ml/min flow rate. Field stimulation was delivered at different rates (0.2–2 Hz) via immersed platinum electrodes running along each side of the perfusion chamber. Simultaneous measurement of stimulation-evoked Ca2+ transients and cell shortening, as well as spontaneous Ca2+ sparks occurring during diastole (16), was acquired in line-scan mode using a 1.3 NA x40 water-immersion objective. A single cell was scanned repetitively (521 Hz) along a line parallel to its longitudinal axis (avoiding nuclei and other high-fluorescence intensity "hot" spots) and extending slightly off cell on both ends. These time-series scans typically were acquired at a pixel size of ~0.05 µm2 x ~1-µm depth resolution. Fluo-4 fluorescence excitation was induced by 488-nm light from the argon laser, with the corresponding fluorescence emission collected by photo-multiplier tube through a 520-nm long-pass filter. Data acquisition was initiated no sooner than 5 min after stimulation had begun to ensure steady-state measurements. To minimize photobleach rundown, laser power was limited to ≤3% of output transmission. In the case of control versus acute application of EtOH trials, minor photobleach rundown was corrected for off-line via interpolation of any decrease in ratio of peak to resting fluorescence (F/F0) between control and washout. In the case of CNTRL versus (chronic) ETOH acquisitions, cells used from each group were subjected to equal numbers and times of scans to minimize photobleach-rundown error.

To provide for the determination of SR Ca2+ content and fractional release, the field stimulation was stopped for a period not greater than 2 s, and immediately thereafter, a short application of Na+- and Ca2+-free (Li+ substituted) Tyrode solution containing 10 mM caffeine was delivered via a multiline-micromanifold fast solution-application device (Warner, CellMicro, ValveLink) so as to evoke the release of total stored SR Ca2+ as measured by peak fluo-4 fluorescence.

For image acquisition under conditions of acute EtOH exposure, Tyrode solution containing either 0.15%, 0.3%, or 1.5% (wt/vol) EtOH was delivered to CNTRL cells for ~10 min per stimulation period via the fast application device. The 0.15% and 0.3% concentrations of EtOH were chosen because they encompass the range of BACs achieved in vivo found for the 1-mo-ETOH and 3-mo-ETOH rats (see above), whereas the 1.5% EtOH represents a near lethal dose of EtOH. For image acquisitions under conditions of chronic EtOH exposure, 1-mo-ETOH and 3-mo-ETOH cells were superfused constantly with Tyrode solution containing 0.15% and 0.19% EtOH, respectively.

Data Analysis

Acquired images were quantitatively analyzed off-line using an IDL software routine (Research Design International). Uncalibrated fluorescence ratios of the line-scan images were analyzed after Gaussian filtering (–3-dB cutoff frequency of 20 Hz) and background subtraction to determine F0 and F. Cell shortening was measured as the percent change in cell length compared with resting cell length (fractional shortening) as determined from the loss of fluorescence at the ends of the line-scan image immediately after the stimulation-evoked Ca2+ transient. The greatest line-mean F/F0 was taken as the Ca2+ transient peak amplitude. The greatest change of fluorescence over time (dF/dt) from start (stimulus) to peak was taken as the Ca2+ transient maximum rate of rise. The reciprocal of the exponential decay time constant ({tau}) fit to the Ca2+ transient from 90% post-peak amplitude to end (next stimulus) was taken as the Ca2+ transient rate of decay. The index of SR Ca2+ content was taken as the peak fluo-4 fluorescence signal resulting from the application of 10 mM caffeine, which was then used to normalize fractional SR Ca2+ release for individual transients.

Analysis of Ca2+ sparks was performed on the same cells in which analyses of Ca2+ transients were performed. Determination of diastolic spark peak amplitude (F/F0), full width (in µm) at half maximum amplitude (FWHM), and full duration at half maximum (FDHM) was accomplished via an automated subroutine based on IDL software and adapted routines (6). Regions of nonregenerative local changes in fluorescence intensity were initially identified as probable Ca2+ sparks and subjected to background subtraction and boxcar filtering over 3–5 pixels (smoothing). The retained smoothed region(s) were then designated as Ca2+ sparks only if the corresponding fluorescence remained elevated for more than 4 ms, if the corresponding decay recovered to levels below 50% of peak in the area of measurement and if there was only one fluorescence peak in the corresponding region of measurement. Spark frequency was determined by calculating the number of conclusively identified Ca2+ sparks that occurred per second of line-scan period per 100 µm line-scan length.

Statistics

Numerical results are expressed as means ± SE, except in Table 1 where values are means ± SD. In determinations regarding acute EtOH effects, paired Student's t-test and/or Bonferroni's one-way ANOVA test was used to assess the statistical differences in results obtained from CNTRL cells before and after acute EtOH application. In determinations regarding chronic EtOH effects, Bonferroni's one-way ANOVA test was used to assess the statistical differences in results obtained from cells isolated from CNTRL and 1-mo-ETOH and 3-mo-ETOH rats. Statistical calculations were made with the use of SigmaStat 2.0 (Systat Software). Differences with P < 0.05 were considered significant.


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  		Table 1. CNTRL vs. ETOH rat average body weight and echocardiography parameters

 

   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Acute EtOH

EtOH effects on cell shortening, Ca2+ transients, and sparks. Whereas the effects of acute EtOH exposure on cell shortening and Ca2+ transients have been described before (7, 8, 52), discrepancies in effective EtOH concentration are apparent, and determinations have not been studied by use of a confocal microscope to examine local fluctuations in intracellular calcium. Thus we were compelled to establish this base action of EtOH on cellular EC coupling and to further such study by examining the action of EtOH on Ca2+ sparks. Figure 1 shows a representative series of Ca2+ fluorescence image recordings of a CNTRL ventricular cardiac myocyte stimulated at 0.5 Hz before and after the addition (10–20 min) of 0.15%, 0.3%, and 1.5% EtOH. Figure 1A illustrates that both cell shortening and Ca2+ transients (quantified via whole cell mean F/F0 intensity vs. time profiles, which are shown above each image) were diminished upon increasing EtOH concentration. This action was reversed after a 5- to 10-min washout of EtOH (data not shown). In the enlarged and enhanced images of Fig. 1B, the dose-dependent decrease in cell shortening induced by EtOH is more clear, as are the incidences of Ca2+ sparks occurring spontaneously during diastole in our isolated cardiac myocytes. A progressive decrease in spark amplitude upon increasing EtOH concentration was suggested in this particular image series. However, statistical analyses of spark incidence (480–277 sparks in 21–32 cells from 4–5 rats) and character in multiple cells indicated that acute exposure to 0.15–1.5% EtOH caused no significant alternations in spark frequency (control vs. 1.5% EtOH, 1.5 ± 0.3 vs. 1.7 ± 0.4 number of sparks·s–1·100 µm–1), amplitude (control vs. 1.5% EtOH, 1.58 ± 0.02 vs. 1.54 ± 0.02 F/F0), width (control vs. 1.5% EtOH: 1.78 ± 0.03 vs. 1.75 ± 0.04 µm at FWHM), or duration (control vs. 1.5% EtOH, 35 ± 1 vs. 33 ± 1 ms at FDHM).


Figure 1
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  		Fig. 1. A: series of ratio of peak to resting fluorescence (F/F0) vs. time confocal Ca2+-fluorescent microscopy line-scan pseudocolor images of cell contractions (white arrowheads signify points of maximal cell shortening) and Ca2+ transients (red line above each image is cellular mean transient) evoked by 0.5-Hz field stimulation of a cell isolated from a control-fed (CNTRL) rat under conditions of control, 0.15%, 0.3%, and 1.5% (wt/vol) ethanol (EtOH). B: same cell as in A but at 0.2-Hz stimulation. Although peak Ca2+ transient intensities are saturated in these enlarged and brightness/contrast-enhanced images (note change in F/F0 calibration bar, denoted by E-F/F0), the incidences of Ca2+ sparks are better evidenced, as are the EtOH dose-dependent decreases in cell shortening.

 
Summarized in Fig. 2, statistical analyses of cell shortening and Ca2+ transient character in multiple cells (32–45 cells from 4–5 rats; stimulation rate, 0.2–0.5 Hz) confirmed that both cell shortening and Ca2+ transient rate of rise were significantly decreased at all EtOH concentrations tested in a dose-dependent manner. Significant decreases in Ca2+ transient peak amplitude and Ca2+ SR content were also evident at EtOH concentrations ≥0.3% and 1.5%, respectively. The rate of Ca2+ transient decay and fractional SR Ca2+ release were not significantly changed by acute EtOH exposure. Results adhered to a similar trend at 1–2 Hz stimulation rates and, when taken together with those at 0.2–0.5 Hz, indicated no significant dependence of these Ca2+ transient parameters on stimulation rate. Thus results at 1–2 Hz are not included here for purposes of presentation clarity and comparison accordance with our Ca2+ spark analyses, the latter of which was conducted only for 0.2- to 0.5-Hz rates where sufficient numbers of sparks can be detected to perform reliable statistics.


Figure 2
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  		Fig. 2. Summary of effects that acutely applied 0.15%, 0.3%, and 1.5% (wt/vol) EtOH exerts on cell shortening (A), Ca2+ transient peak amplitude (B), Ca2+ transient maximal (Max) rate of rise [maximal change of fluorescence over time (dF/dt); C], Ca2+ transient rate of decay (reciprocal of {tau} exponential time constant; D), sarcoplasmic reticulum (SR) Ca2+ content (E), and SR Ca2+ fractional release (F) in cells isolated from CNTRL rats. Ca2+ transients were evoked by 0.2- to 0.5-Hz field stimulation, from which SR Ca2+ content and fractional release were determined from peak amplitudes of transients evoked by 10 mM caffeine (0.2–0.5 Hz prestimulated). *P < 0.05, **P < 0.01, ***P < 0.001.

 
Chronic EtOH

Demographic and echocardiographic parameters in ETOH versus CNTRL rats. Body weights, left ventricular mass and left ventricular mass-to-body weight, and echocardiographic parameters (Table 1) were not significantly different between 1-mo-ETOH (n = 6) and 1-mo-CNTRL rats (n = 6) or between 3-mo-ETOH (n = 6) and 3-mo-CNTRL rats (n = 6). Collectively, these data indicate, similar to our previous finding (23), that <4 mo of heavy ethanol consumption is not associated with the development of dilated cardiomyopathy in Sprague-Dawley male rats.

Cell shortening, Ca2+ transients, and sparks in ETOH versus CNTRL cardiac myocytes. Figure 3 shows Ca2+ fluorescence image recordings of 0.3-Hz field-stimulated ventricular cardiac myocytes isolated from 1-mo-CNTRL rats versus 1-mo-ETOH rats (Fig. 3A) and from cells isolated from 3-mo-CNTRL rats versus 3-mo-ETOH rats (Fig. 3B). The recordings show that the contractions, Ca2+ transients, and also Ca2+ sparks in cells from 1-mo-ETOH rats are greater than those in controls. In contrast, these characteristics of EC coupling were reduced in cells from 3-mo-ETOH rats versus controls.


Figure 3
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  		Fig. 3. A: real-time confocal Ca2+-fluorescent microscopy line-scan pseudocolor images (F/F0 vs. time) of cell contractions and Ca2+ transients evoked by 0.3-Hz field stimulation in a cell isolated from a 1-mo-CNTRL rat vs. that from 1-mon EtOH-fed (1-mo-ETOH) rat. White double arrows indicate points of maximal cell shortening. Lower image in each is brightness/contrast-enhanced (refer to lower E-F/F0 calibration bar) to better illustrate Ca2+ sparks occurring during stimulus cycles. B: as in A but in a cell isolated from a 3-mo-CNTRL rat vs. that from a 3-mo-ETOH rat.

 
Analyses of cell shortening and Ca2+ transients in cardiac myocytes from 1-mo-ETOH (25 cells from 4 rats) versus CNTRL rats (20 cells from 4 rats), as illustrated in Fig. 4, indicated that cell shortening, Ca2+ transient peak amplitude, and rate of rise along with SR Ca2+ content were all increased in cells from 1-mo-ETOH rats versus controls. In contrast, these EC coupling parameters were all decreased in cells from 3-mo-ETOH rats (18 cells from 4 rats) versus 3-mo-CNTRL rats (15 cells from 3 rats). There was a tendency toward increases in Ca2+ transient rate of decay and fractional SR Ca2+ release in cells from 1-mo-ETOH rats vs. CNTRL rats that did not reach statistical significance. However, there was a decrease in the rate of Ca2+ transient decay in cells from 3-mo-ETOH rats versus CNTRL rats that did reach statistical significance.


Figure 4
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  		Fig. 4. Summary comparing cell shortening (A), Ca2+ transient peak amplitude (B), Ca2+ transient maximal rate of rise (maximal dF/dt; C), Ca2+ transient rate of decay (reciprocal of exponential time constant {tau}; D), SR Ca2+ content (E), and fractional SR Ca2+ release (F) in cells isolated from 1-mo (or 1m)-CNTRL vs. 1-mo-ETOH rats and 3-mo (or 3m)-CNTRL vs. 3-mo-ETOH rats. Ca2+ transients were evoked by 0.3-Hz field stimulation, from which SR Ca2+ content and fractional release were determined from peak amplitudes of transients evoked by 10 mM caffeine (0.3 Hz prestimulated). *P < 0.05 and **P < 0.01.

 
Following a similar trend, statistical analyses of Ca2+ sparks, summarized in Fig. 5, indicated that sparks in cardiac myocytes from 1-mo-ETOH rats (302 sparks in 25 cells from 4 rats) occurred at the same frequency, but their peak amplitude and widths were significantly increased compared with sparks in 1-mo-CNTRL rats (325 sparks in 20 cells from 4 rats). In contrast, sparks in cells from 3-mo-ETOH rats (344 sparks in 18 cells from 4 rats) occurred at greater frequency with decreased peak amplitudes and widths compared with sparks in cells from 3-mo-CNTRL rats (243 sparks in 15 cells from 3 rats). No significant difference in spark duration was detected between groups.


Figure 5
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  		Fig. 5. Summary comparing Ca2+ spark frequency (A), peak amplitude (B), width (C), and duration (D) of Ca2+ sparks occurring spontaneously between 0.3-Hz field stimuli in cells isolated from 1-mo-CNTRL vs. 1-mo-ETOH rats and 3-mo-CNTRL vs. 3-mo-ETOH rats. **P < 0.01 and ***P < 0.001.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The goal of the present study was to provide insight into how the acute negative inotropic effects of acute EtOH exposure are translated to the chronic EtOH condition by detailing how various characteristics of cellular EC coupling in isolated rat cardiac myocytes change upon acute (10–20 min) and early chronic (1–3 mo) EtOH exposure. The results of our investigation are collectively recompiled in Table 2 and presented as percent change for each EtOH administration condition relative to the respective controls. This overall comparison makes it clear that the negative inotropic action and associated defects in EC coupling induced by acute EtOH exposure are not reflected at 1-mo chronic EtOH exposure, and, in fact, the data clearly demonstrate positive inotropy and associated enhancement in EC coupling at this very early stage in chronic EtOH exposure. However, at 3 mo chronic EtOH exposure, there is a recurrence of negative inotropy and aberrant EC coupling similar but not identical to that seen upon acute exposure to high concentrations of EtOH. These experiments provide the first evidence for distinct biphasic effects of chronic EtOH on cardiac cellular function.


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  		Table 2. Acute vs. early chronic EtOH exposure on EC coupling on rat ventricular cardiac myocytes

 
Negative Inotropic Effects of Acute EtOH Exposure in Cellular EC Coupling

Our first series of experiments was intended both to establish a comparison of our findings that employed our particular methodology and conditions with previous studies documenting the negative inotropic effects of acute EtOH exposure and to newly document the effects of EtOH directly on individual Ca2+ release units as assessed by measurement of changes in the characteristics of Ca2+ sparks. We did find that acute application of EtOH ≤0.15% reduced cell shortening, which commenced before reductions of Ca2+ transient amplitude at EtOH ≤0.3%. Although these results were similar in action to that of previous findings (7, 52), we found EtOH to be somewhat more potent than reported in those studies, perhaps owing to the greater detail afforded by confocal determinations. Significant reductions in SR Ca2+ content and Ca2+ transient decay rate were also evident but only at acutely lethal concentrations of EtOH. In any event, our results still concur with the proposed mechanism of EtOH acting at lower concentrations to interfere with myofilament cross-bridge formation and thus decreasing myofilament sensitivity to Ca2+ released from the SR, which is then exaggerated by reduced global SR Ca2+ release that occurs at higher EtOH concentrations (7, 14, 15, 20). Further contributing to these negative inotropic effects is attenuation in cardiac L-type Ca2+ channel current (ICa-L) reported by our group (3) and others (18, 35), which constitutes an EtOH-induced reduction in EC coupling trigger Ca2+.

As for acute EtOH actions on individual SR Ca2+ release units, our results suggest no significant changes in Ca2+ spark character under our experimental conditions. This would seem to indicate that even though acute EtOH exposure reduces the ICa-L trigger Ca2+, it is evidently still sufficient to activate SR ryanodine receptor Ca2+ release channels (RyRs). Moreover, this also seems indicative that RyR function is evidently not significantly altered by acute EtOH exposure. This issue perhaps warrants further investigation given that earlier reports regarding EtOH effects on spontaneous SR Ca2+ release are conflicting, with both enhancement and inhibition being observed (36, 40). What may be apparent in all of this is that small EtOH effects on various multiple components governing global Ca2+ release combine to diminish Ca2+ transients.

Early Effects of Chronic EtOH Exposure on Cellular EC Coupling

Surprisingly, and in contrast to the negative inotropic effects observed in acute EtOH exposure, we found that even though the rats had attained BACs of ~0.15% EtOH after 1 mo of chronic EtOH consumption, cardiac myocytes from these 1-mo-ETOH rats exhibited increased cell shortening, Ca2+ transient magnitude and rate of rise, as well as increased in spark magnitude and width. Thus 1 mo of chronic EtOH exposure actually promotes positive inotropy. This effect is not attributable to acute EtOH withdrawal because we maintained the cells in 0.15% EtOH throughout their isolation and all subsequent studies. Hence, these positive inotropic effects most likely stem from the increased SR Ca2+ content, which would explain the increased Ca2+ transient amplitude signifying greater global SR release, as well as the increased Ca2+ spark amplitude signifying greater SR release at the level of individual Ca2+ release units. The increase in the total amount of Ca2+ released for each spark may have also allowed for greater spread between release units indicated by the increased spark width and the more efficient EC coupling. Note that there was no change in spark frequency, thus no associated increased SR Ca2+ leak and reduced load. The increase in the efficiency of EC coupling occurred without an increase in the fractional release of Ca2+ during the SR, so that this particular positive inotropic effect was almost entirely related to the increase in SR Ca2+ content.

After an additional 2 mo of chronic EtOH exposure, virtually each component of EC coupling that had been enhanced at 1 mo of chronic EtOH was then reduced. Thus a negative inotropic effect reminiscent of acute EtOH exposure was manifest at 3 mo of chronic EtOH. There were, however, clear differences in the altered EC coupling at 3 mo of chronic EtOH exposure and that induced by acute EtOH exposure, the most prominent of which was increased Ca2+ spark frequency that was accompanied by decreased spark amplitude and width. The increased spark frequency likely corresponds to a reported increase in SR Ca2+ leak in alcoholic rats (39) and may be indicative of a qualitative change in RyR behavior that has been associated with hyperphosphorylation of RyRs in a number of types of heart failure (30). The latter remains to be verified, but the resultant increased SR Ca2+ leak upon chronic EtOH exposure would contribute to decreased SR Ca2+ load, which in turn would result in smaller Ca2+ transients that rise and decay more slowly, which consequently would diminish cell shortening. As with the positive inotropic effects observed after 1 mo, these changes in EC coupling took place in the absence of any changes in fractional SR Ca2+ release, indicating that the decrease in SR load seems to be a key means by which these 3-mo chronic EtOH negative inotropic effects occur in this model.

The basis for the transitory increase in SR Ca2+ content at 1 mo and its subsequent decrease at 3 mo of chronic EtOH exposure is not clear from our results. However, previous reports regarding chronic EtOH effects on sarco(endo)plasmic reticulum Ca2+-ATPases (SERCA) collectively suggest similar transitory changes. It has been reported that increased skeletal muscle (SERCA1) protein expression and ATPase activity occur after 6 wk of chronic EtOH exposure (38), and this is accompanied by a reported increase in cardiac SERCA2 activity after 10 wk of chronic EtOH exposure (17). Conversely, SERCA2 protein expression has been reported to be unchanged after 7 mo of chronic EtOH exposure (11), and SR Ca2+ binding and uptake have been reported to be inhibited after 3 mo of chronic EtOH exposure (41). Thus early chronic EtOH appears to upregulate SERCA function and/or expression, whereas extended chronic EtOH appears to downregulate it. However, for this contention to hold, further study is required to ensure comparability between SERCA1, SERCA2, and the SERCA regulatory protein phospholamban (not to mention species).

Additionally, chronic EtOH-induced changes in intracellular Na+ concentration ([Na+]i) can alter SR Ca2+ content. Both acute and chronic EtOH exposure are known to induce an increase in sarcolemmal Na+ permeability by ambiguous mechanism(s), which results in an elevated [Na+]i early (1–2 mo) in chronic EtOH exposure that returns to near-normal levels later (+4 mo) at the expense of increased Na+/K+ ATPase activity and/or protein expression (5, 10, 21). An accumulation of intracellular Na+ could indeed cause an increase in total intracellular Ca2+ concentration ([Ca2+]i) and thus presumably SR Ca2+ load via reverse-mode Na+/Ca2+ exchanger (NCX) activity, given no chronic NXC inhibition (33). This would also effectively counter both the reduced myofilament Ca2+ sensitivity and ICa-L-based trigger Ca2+. If [Na+]i and [Ca2+]i returned to near-normal levels later in chronic EtOH exposure, the normal but lower [Ca2+]i would no longer compensate for reduced myofilament Ca2+ sensitivity and ICa-L trigger Ca2+. Although such a scenario remains to be and could be tested, this along with possible temporal regulation of SERCA as mentioned above could give rise to an initial phase of positive inotropy that is followed by negative inotropy during early chronic EtOH exposure. In fact, the involvement of [Na+]i in these events may be inferred from our observation that both peak and window rapid Na+ current (INa) increase at 1 mo of chronic EtOH exposure, whereas they return back to baseline levels at 3 mo of chronic EtOH exposure (3). In contrast, ICa-L was reduced throughout chronic EtOH exposure. Such changes in INa during chronic EtOH exposure not only seem to closely parallel those for [Na+]i, but the increased window INa may also constitute an early component of the as yet unidentified EtOH-induced sarcolemmal Na+ leak.

Acute Negative Inotropic Effects of EtOH Exposure Do Not Immediately Translate Early in Chronic EtOH Exposure

Interestingly, the acute negative inotropic effects of EtOH were not found after 1 mo of EtOH consumption. Rather, we found a positive inotropic phase and enhanced EC coupling. This positive inotropic effect or phase may reflect the development of tolerance, which serves as a mechanism that allows the heart to counteract the acute negative effects of EtOH. The 1-mo positive inotropic phase is followed at 3 mo by attenuated EC coupling and a negative inotropic effect. This 3-mo effect may represent an important change in EC coupling and intrinsic myocyte function that leads to the development of ACM. It is important to note that both the 1- and 3-mo phases were not associated with EtOH-induced myocardial structural changes because we found no significant changes in echocardiographic parameters between EtOH and control groups. Thus, key to a primary goal of this study to examine EC coupling changes at an early stage of ACM, our findings here may be indicative of causal rather than consequential events in the development of cardiovascular diseases induced by chronic EtOH, such as, but not limited to, ACM. Indeed, we have previously shown in this model that a longer period of EtOH consumption is associated with the development of ACM, which thus lends support for the use of this model to investigate early and late EtOH-induced changes in the myocardium. It remains to be determined whether longer periods of EtOH consumption are associated with further and persistent changes in EC coupling.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
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REFERENCES
 
This study was supported by National Institute on Alcohol Abuse and Alcoholism Grants R01-AA-10969 and R21-AA-013915.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. Aistrup, Dept. of Molecular Pharmacology & Biological Chemistry, Northwestern Univ., Feinberg School of Medicine, 303 E. Chicago Ave., Chicago, IL 60611 (e-mail: glais{at}northwestern.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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Ethanol and cardiac function
Am J Physiol Heart Circ Physiol, September 1, 2006; 291(3): H1027 - H1028.
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