Endurance exercise training increases the functional capacity of cardiovascular system. The adaptations include improved fractional shortening as well as increases in cardiac output, stroke work, cardiac mass, and dimensions (14,28). Although many of these adaptations have been delineated at the physiological and morphological levels, very little is known about the underlying molecular changes that are responsible. Some training studies have demonstrated changes at the mRNA level for different functionally important molecules including cardiac troponin-I, atrial natriuretic factor (ANF), β-creatine kinase, and skeletal α-actin genes (4,20). Although it is likely that exercise-induced shifts in the skeletal muscle phenotype are regulated at the translational and transcriptional levels, very little is known about the phenotypic shifts that occur in the myocardium in response to exercise (22).
Cyclosporin A (CsA) is an immunosuppressive drug that has seen great use in transplant programs. Unfortunately, CsA is thought to be at least partially responsible for the decreased exercise capacity observed in transplant patients and in chronic-exercise animal studies (25,35). Other negative side effects of CsA include nephrotoxicity, hepatotoxicity, and hypertension. Additionally, CsA has been shown to alter mitochondrial function in several organs including skeletal muscle (1,15). CsA binds to the intracellular cyclophilin receptors and inhibit calcium-activated protein phosphatase 2B, also known as calcineurin (23). Calcineurin is a key enzyme involved in the regulation of calcium-dependent pathways.
A significant role for the calcium dependent pathways in the myocardium was established by Sussman et al. (31). This group demonstrated that CsA blocked pressure overload-induced cardiac hypertrophy. This interference indicates a role for calcium dependent pathways in regulating cardiac hypertrophy and modifying cardiac gene expression in response to overload. It is unclear whether these same pathways are critical in the response to exercise-induced cardiac growth. Exercise training has been shown to counteract some of the adaptations associated with pathological overload (27,28). It is thought that training induces a “balanced growth” of cardiomyocytes as suggested by the maintenance of the myofibril:mitochondrial ratio (32,33).
We are interested in studying the early responses to exercise-training–induced overload because in this phase the heart has not compensated for the increased functional demand and has not undergone significant hypertrophy. It is during this period that the imbalance between increased demand and cardiac functional capacity is greatest. And it is likely that at this time, the mechanisms responsible for propagating changes in the myocardial phenotype are most active. We have previously studied early training-induced changes in cardiac growth and ANF gene expression (6,7). It is unclear whether the calcium dependent pathways that are critical to pressure-overload–induced hypertrophy have a significant role in the early adaptations to exercise training. Exercise and pressure overload are remarkably different forms of functional overload. We hypothesized that cyclosporin would not alter exercise-training–induced cardiac growth. To investigate this, we have exercise-trained animals that were injected with either CsA or vehicle for 1 wk.
Cyclosporin A was obtained from Calbiochem (La Jolla, CA). The antimyosin antibody was purchased from Chemicon (Temecula, CA). The secondary antibody used was part of the Amersham ECL Plus kit (Amersham, Piscataway, NJ). Restriction enzymes, T4 DNA ligase, and other enzymes were purchased from either the Promega Corporation (Madison, WI) or New England Biolabs (Beverly, MA). 32P-labeled nucleotides were purchased from New England Nuclear (Boston, MA). Other reagents were purchased from Fisher Scientific, Sigma-Aldrich, or as noted.
Animals and exercise training protocol.
Twenty male Sprague-Dawley rats were utilized and randomly assigned to sedentary or exercise-training groups. These groups were further subdivided to vehicle injection (50% ethanol) or CsA injection (15 mg·kg−1·d−1) groups. Injections were i.p., between 0900 and 1000 h, and those animals in the exercise-training group were swum between 1400 and 1600 h. The animals were swum three to five per tank, with a surface area of 785 cm2 and with the water temperature being maintained between 31 and 35°C. Before the training period, the animals were familiarized with the water by placing them in shallow water and then in the swimming tank to swim briefly. For the exercise-training protocol, animals swam 10–20 min the first day and then 60 min·d−1 for the next 6 d. Animals were closely monitored to ensure animal safety. Animals were also monitored to ensure that they did not spend time underwater holding their breath and potentially creating a hypoxic stimulus. After the swimming session, the animals were dried before being returned to their cages. Animals were sacrificed on the day after their final training bout, by injection of pentobarbital (75 mg·kg, i.p.). An additional 10 animals were used to determine the effects of CsA on βMHC protein levels. The effectiveness of the CsA injections was determined by TDX analysis (Abbot Laboratories; Chicago, IL) of blood samples taken at the time of sacrifice and 20–24 h after the last exercise bout. Cyclosporin-injected animals had blood levels greater than 1500 mg·mL−1 compared with vehicle, which were less that 25 ng·mL−1. Animals were housed in the Department of Comparative Medicine, maintained on a standard diet, and on a 12-h light/dark cycle. Experimental protocols had institutional approval and followed ACSM guidelines. Animals were maintained in accordance the Guide for the Care and Use of Laboratory Animals as described in the Animal Welfare Act (PL-94-279).
At the time of sacrifice, the animals were weighed to determine body weight (BW) and the hearts removed, washed briefly in ice-cold phosphate-buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.4 mM KH2PO4, pH 7.3), and blotted. Hearts were weighed and then separated by chamber and individual chamber weights were recorded. The left ventricle (LV) was divided to separate the endocardial and epicardial layers. Tissues were then frozen in liquid N2 and stored at −80°C until prepared for analysis. RNA was prepared using a modification of the method described by Chomczynski and Sacchi (5). RNA was quantified by spectrophotometry (40 μg/1 O.D. at 260 nm). RNA was blotted on Hybond-N+ membrane (Amersham, Piscataway, NJ) and fixed by UV cross-linking. Total RNA was analyzed by slot blot analysis, probing first with a gene-specific probe and then with a 28S-ribosomal probe, the latter being used to normalize for loading variations. Membranes were prehybridized for 4–6 h at 42°C, in hybridization buffer (5x SSC, 5x Denhardt’s, 1% SDS, 0.2 mg·mL−1 sonicated salmon sperm DNA, and 50% formamide) before labeled probe (106 cpm·mL−1) was added. Hybridizations were carried out overnight at 42°C. The separate probes used included: 1) rat ANF cDNA (30); 2) ribosomal 28S coding oligonucleotide (Clonetech, Palo Alto CA); and 3) single-strand oligonucleotides coding for either the αMHC or βMHC 3′UTR regions. Double-strand cDNA probes were labeled by a random primer reaction using α-32P-dCTP; single-strand DNA was labeled using T4 polynucleotide kinase and γ-32P-ATP. After hybridizations, blots were washed twice at 27°C in 2xSSC, 1% SDS and twice at 27°C in 1xSSC, 1% SDS. The progress of the wash was monitored by a Geiger counter. after exposure to the ANF, αMHC, or βMHC probes, the blots were stripped and reprobed using a 28S-ribosomal oligonucleotide probe. Band intensities were quantified by phosphoimage analysis by using a Storm 840 PhosphoImager (Molecular Dynamics, Palo Alto CA). The RNA data reported are the ratio of the gene-specific band to 28S-band intensity.
Western blot analysis.
Left ventricular extracts were homogenized in ice-cold PBS buffer. Protein concentration was determined by the Bradford method (BioRad, Hercules, CA); 30-μg protein was loaded on an 8% SDS-PAGE gel, and electrophoresis was carried out for 2 h before the proteins were transferred to a Hybond-P membrane (Amersham). The membranes were blocked using a 5% nonfat milk in PBS overnight at 4°C. The media was exchanged and a 1:1000 dilution of the monoclonal anti-β myosin antibody was added for 16 h at 4°C. The blots were washed briefly and a 1:4000 dilution of the anti-mouse peroxidase-labeled secondary antibody was added; the blots then incubated for 1 additional hour at room temperature before being washed. Antibody binding was visualized using the Amersham ECL Plus kit (Amersham), and band density was quantified from a digitized picture by using AlphaEase software (Alpha Innotech, San Leandro, CA).
Cytochrome oxidase assay.
Cytochrome oxidase activity was determined using a modified method of Wharton and Tzagoloff (34). In brief, plantaris muscle was homogenized in ice-cold buffer (250 mM sucrose, 100 mM KPO4, pH 7.0) before being centrifuged at 2000 ×g to remove debris. The supernatant was retained and protein concentration determined. A constant amount of protein was added to a buffered solution of reduced cytochrome C (10 mM KPO4, pH 7.0, 0.07% reduced cytochrome C). The rate of cytochrome c oxidation was followed at 550 nm (ςε = 21.0 mM−1·cm−1), using a Beckman model 35 spectrophotometer (Beckman Instruments, Fullerton, CA).
Statistical comparisons were made using a two-way or three-way ANOVA (P < 0.05) as appropriate. Post hoc analysis was performed using a Fisher’s LSD multiple-comparison test. Statistical analyses were performed using NCSS 2000 software (NCSS, Kaysville, UT)
Endurance exercise training is characterized by an increase in aerobic capacity. Because we were unable to determine an animal’s V̇O2max, a biochemical marker of skeletal muscle respiratory capacity was utilized. Exercise-induced increases in mitochondrial enzyme activity are a hallmark of endurance training (16). One week of exercise training caused a significant increase in plantaris muscle cytochrome oxidase activity (Fig. 1), indicating an aerobic adaptation to exercise. Separate from the effects of exercise, CsA also induced a significant increase in cytochrome oxidase activity in the sedentary animals. As will be discussed below, this may be the result of CsA’s direct effect on the mitochondria.
The LV/BW ratio of the exercise trained animals was significantly increased by 11.9% over that of the sedentary animals, and similar increases were observed in both the vehicle as well as the CsA-injected animals (Fig. 2B). There were no significant differences in the RV weights or RV/BW of the control and exercise-trained animals. The small increases in the LV/BW ratio are consistent with what we have demonstrated previously using the 1-wk swim-training protocol (6,7). Although this index is useful in experimental paradigms where body weight is unaltered, exercise tends to decrease the rate of growth of male rats. We have previously used the swim-training protocol to demonstrate that in female animals, the LV/BW ratio was increased by training, without a significant change in BW (7). In the present study, differences in body weight were influenced more by drug treatment than by exercise (Table 1). Two-way ANOVA indicated that drug treatment but not exercise significantly altered body weight. However, when LV weight (Fig. 2A) was plotted as a function of body weight, exercise training produced a significant increase in LV mass, irrespective of drug treatment. These results indicated that 1 wk of swim-training protocol produced a similar degree of hypertrophy in both vehicle- and CsA-treated animals.
Consistent with our previous studies, left ventricular ANF-mRNA expression levels in sedentary animals were very low. One week of exercise training was accompanied by a significant increase in the ANF-mRNA levels (Fig. 3). Although the ANF-mRNA levels appear to be higher in the trained CsA-treated animals compared with the trained vehicle, they were not statistically different. At the time of sacrifice, the left ventricular samples were divided to separate the endocardial and epicardial layers. No differences in transmural ANF expression were observed (data not shown).
Previous studies have found an exercise-induced increase in myofibrillar ATPase activity (27). This increase in ATPase activity may be attributed to an increase in the V1 isoform that is the αMHC gene product. Although swim training of the vehicle-injected animals was associated with a 45% rise in αMHC-mRNA, these values were not found to be significantly different. It was observed that βMHC-mRNA levels were increased in all CsA-injected animals, whereas training did not influence βMHC-mRNA levels (Fig. 4).
To confirm that the CsA-induced changes in βMHC expression were reflected at the protein level, a βMHC-specific antibody was used to probe left ventricular protein extracts from vehicle- or CsA-treated animals. In agreement with the RNA data, we observed a five-fold increase in βMHC protein (Fig. 5).
We have utilized a 1-wk swimming program to determine whether CsA will influence exercise-induced cardiac hypertrophy. In both vehicle- and CsA-injected animals, exercise training induced cardiac growth. These results indicate that the CsA-sensitive calcium-dependent pathways were not critical for exercise-induced cardiac growth. These findings are remarkably different from some pathological forms of overload-induced hypertrophy in which calcium-dependent signal pathways are important.
CsA is an immunosuppressive drug commonly utilized in transplant programs. CsA is of interest in the study of cardiac hypertrophy due to its inhibitory effect on calcineurin, a key phosphatase that mediates calcium-dependent events within the myocardium. However, CsA is somewhat cardiotoxic and adversely effects cardiac performance. Kingma et al. (18) demonstrated that peak systolic pressures were significantly lower in CsA-treated animals. Banijamali et al. (2) found that CsA treatment depressed myocardial contractility. It has been suggested that CsA alters calcium release from the sarcoplasmic reticulum and that chronic drug treatment results in a net depletion of calcium stores (2). Beyond that effect, CsA also alters the functional properties of mitochondria. CsA treatment of isolated hepatocytes significantly increased cytochrome oxidase activity and carnitine acetyl transferase but also decreased the respiration rate of gastrocnemius muscle (1,15). Collectively, these studies suggest that CsA can influence the functional capacity of the heart by a direct effect on the intracellular processes.
That CsA did not block exercise-induced cardiac growth stands in contrast to that found by Sussman et al. (31), who demonstrated that CsA blocked pressure-overload–induced cardiac hypertrophy. These findings suggest that separate signal transduction pathways were activated for each form of overload. There are several differences between these forms of overload. First, exercise induces a mild form of volume overload that promotes ventricular growth patterns that differ from pressure overload. Second, most models of pressure overload maintain a constant increase in overload. In contrast, exercise applies an overload only for a short period in the total sum of the day. This permits the heart to recover from the stress. This point may be important since the chronic overload may lead to the recruitment of calcium-activated proteases, which have been hypothesized to be responsible for depressed myocardial contractility in failing hearts (12). Third, in the model used by Sussman et al. (31), pressure overload was induced by stenosis of the superior abdominal aorta, which has the immediate effect of decreasing renal blood flow. This decrease leads to an activation of the renin-angiotensin system, a strong stimulus for cardiac hypertrophy. However, a report by Geenen et al. (13) has ruled out a significant role for angiotensin in exercise-induced cardiac growth. Further, our own studies have found that the critical cis-elements that control exercise-induced increases of ANF transcription are separate and different from angiotensin activation of ANF transcription (6,8). Collectively, these findings suggest that the signal transduction pathways mediating cardiac growth in response to chronic exercise are different from those recruited in response to pathological forms of cardiac overload.
That CsA did not block exercise-induced cardiac growth is a finding that stands in contrast to that reported by Eto et al. (11). In that study, CsA-injected animals did not demonstrate exercise-induced cardiac growth. However, these two studies are remarkably different. Eto et al. utilized a voluntary model of run training, and it is not possible to determine from that report whether CsA treatment altered the intensity of exercise performed or the total amount of work performed. CsA decreases muscle respiration capacity, and this correlates with decreased endurance capacity in animals (15,25). In contrast, the present study used a forced swim-training protocol. McArdle (21) demonstrated that swimming rats increased their oxygen consumption to more than 60% V̇O2max. An 8-wk swim-training program demonstrated that the trained rats had a lower resting heart rate, increased coronary blood flow, increased heart weight, and greater functional capacity compared with sedentary animals (29). As indicated in Methods, the animals were made to swim as a group to ensure that exercise intensity was elevated and that all the animals performed the same total workload. Our results are similar to what we have seen previously and to others that have reported small increases in ventricular weights and increased cytochrome C content as an early adaptation to training (6,7,14). It is possible that with voluntary exercise the intensity of exercise and/or volume of exercise were substantially less in the CsA-treated animals, and this may account for the lack of cardiac growth.
We have previously demonstrated that exercise training will increase plantaris cytochrome oxidase activity (7,9,10). An unexpected result was that that the CsA increased plantaris cytochrome oxidase activity in the sedentary animals. Because this parameter has been used as an index of muscle aerobic capacity, the finding is a paradox. Backman et al. (1) reported a similar increase of cytochrome oxidase activity by using cultured hepatocytes or HepG2 cells. Exposure to CsA increased cytochrome oxidase activity 190%, whereas other mitochondrial enzymes were either increased or uninfluenced. Although the mechanism remains unclear, it has been suggested that CsA’s interaction with the mitochondrial membrane Ca+2 pores may be involved, and this may have induced some mitochondrial proteins whereas leaving others unaffected. An alternative explanation may be that CsA induced the synthesis of endothelial nitric oxide synthase, which should elevate nitric oxide (NO) levels (3). NO has been shown to lower mitochondrial respiration by a direct inhibition of the cytochrome oxidase complex, and this may have signaled an increase in cytochrome oxidase synthesis (3). This idea is supported by the findings of Lehrer-Graiwer et al. (19), who demonstrated that NO induced cytochrome oxidase expression. In the present study, although the trained animals receiving CsA had the highest cytochrome oxidase levels, the values were not significantly greater than the vehicle-treated trained animals, suggesting that the effects were not additive.
Concomitant with pressure-overload–induced cardiac growth is a significant shift toward the myocardial embryonic phenotype. It is characterized by an increased ventricular expression of a number of molecules including the ANF, α-skeletal actin, and the βMHC gene products (17). Exercise-induced increases in ANF expression have also been observed by our group and others (4,7,20). It is unclear what the exercise-induced shifts in MHC expression might be. Exercise training will increase the myosin V1 isoform component as well as increase myosin ATPase activity (28,29). We did not find an exercise-related shift in the MHC expression patterns. However, βMHC-mRNA was elevated in all CsA-treated animals, and this shift was also reflected at the protein level. It is unclear whether this was a direct effect of CsA on βMHC transcription or a compensatory mechanism. Meissner et al. (24) demonstrated that CsA blocked increased βMHC expression in electrostimulated cultured muscle cells. This would suggest that the increases in βMHC expression observed might have been compensatory. The CsA-induced shift in the myosin phenotype offers a reasonable explanation for the decreases in myocardial function (18). These two observations, a phenotype shift and depressed cardiac function, indicate a compromised exercise profile and offer an explanation for lower exercise capacity in otherwise healthy transplant patients receiving CsA (35).
Exercise increases the functional capacity of cardiovascular system by a shift in both its morphology and phenotype. Although some of these adaptations have been delineated at the physiological and morphological levels, very little is known about the underlying molecular changes that are responsible. CsA inhibits a key enzyme calcineurin, which is important for the regulation of calcium-dependent signaling pathways in the myocardium, and we have used CsA to determine whether these pathways were important in the early adaptations to exercise training. The present study has two major observations: 1) CsA did not influence exercise-induced cardiac growth, and 2) CsA did induce a phenotype shift in the myocardium. The first finding indicates that exercise-induced cardiac growth utilizes mechanisms that are separate and distinct from those activated by pathological forms of pressure overload. The second finding suggests a compromised phenotype in transplant patients receiving CsA, and this may have an impact on their participation in exercise programs.
This work has been supported in part by NIH: R29HL59417 and P01HL43023.
Address for correspondence: John G. Edwards, Ph.D., Department of Physiology, New York Medical College, Valhalla, NY 10595; E-mail: email@example.com.
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Keywords:©2002The American College of Sports Medicine
TRAINING; MYOCARDIUM; HEART; IMMUNOSUPPRESSION