Skeletal Muscle Sarcoplasmic Reticulum Ca2+-ATPase Gene Expression in Congestive Heart Failure
Abstract Congestive heart failure leads to skeletal muscle abnormalities, one of which is a prolongation of sarcoplasmic reticulum Ca2+ flux. The purpose of this study was to determine whether skeletal muscle of spontaneous hypertensive and heart failure rats have alterations in the expression of the sarcoplasmic (or endoplasmic) reticulum Ca2+-ATPase (SERCA) gene. Northern analysis revealed that SERCA1, the predominant skeletal muscle isoform, was decreased by 45%, 43%, and 58% in the tibialis anterior, plantaris, and diaphragm muscles, respectively. Ribonuclease protection assay showed that the decrease was due to the adult isoform, SERCA1a, with minor changes in the alternatively spliced neonatal isoform, SERCA1b. There was no change in SERCA1 mRNA levels in gastrocnemius muscles. No change was found in SERCA2a (cardiac/slow skeletal isoform) mRNA or protein levels or in SERCA2b (smooth muscle isoform), dihydropyridine receptor, or α-actin mRNA levels in diaphragm muscle. Northern blot and ribonuclease protection assays showed that SERCA2a decreased 61% in the heart while the alternatively spliced isoform, SERCA2b, decreased 27%. Western analysis of the tibialis anterior, diaphragm, and gastrocnemius muscles showed a decrease in SERCA1 protein levels by 46%, 64%, and 42%, respectively, whereas sarcoplasmic reticulum Ca2+-ATPase activity, a functional correlate of SERCA expression, was decreased by 38%, 38%, and 40% in the same muscles. SERCA2 protein expression decreased by 36% in the failing heart. Decreases in both mRNA and protein suggest pretranslational control of SERCA1 expression, whereas the lack of decreased SERCA1 mRNA in gastrocnemius muscle suggests translational regulation. The decreased SERCA1 protein expression in all muscles studied probably contributes to contractile abnormalities related to excitation-contraction coupling function in heart failure.
Skeletal muscle fatigue is a major contributor to work intolerance and morbidity in patients with CHF.1 Skeletal muscle impairments are evident in part because patients treated with medications that improve cardiovascular function, such as ACE inhibitors, still report debilitating muscular fatigue.2 Also, it has long been known that exercise intolerance in CHF is greater than that predicted by the extent of left ventricular dysfunction (reviewed in References 2 and 32 3 ). A major question has been whether skeletal muscle impairments upon exertion are due to physical inactivity secondary to central limitations. This is because the changes are similar to those resulting from deconditioning such as low oxidative capacity, early onset of lactate accumulation, increased fatigability,4 and slow to fast shifts in myosin heavy chain isoform expression.5 6 However, it was recently shown that decreases in oxidative enzyme activity and alterations in myosin heavy chain isoform expression occurred in muscles from CHF rats that had the same physical activity levels as age-matched controls.6
Another functional abnormality in skeletal muscle in CHF is related to the intracellular Ca2+ flux associated with excitation-contraction coupling. Skeletal muscle in rodents with heart failure shows prolonged intracellular Ca2+ transients during twitch and tetanic contractions and accelerated fatigue development,7 slowed rates of tension development and relaxation,8 and slowed SR Ca2+ uptake rate and Ca2+-ATPase activity.9 Again, physical inactivity is unlikely to cause these changes because it has the opposite effect; that is, inactivity leads to faster SR Ca2+ cycling characteristics.10 11 12 Nevertheless, the molecular mechanism of these defects in skeletal muscle is not known, but it could involve alterations in the expression of the major rate-limiting protein of intracellular Ca2+ sequestration, SERCA, as is found in the failing myocardium (reviewed in Reference 1313 ).
There are three known SERCA genes, two of which (SERCA1 and SERCA2) have alternative splicing products. The predominant isoform expressed in skeletal muscle is SERCA1a, and its alternative splicing product, SERCA1b, is expressed in neonatal skeletal muscle.14 15 16 The predominant isoform expressed in cardiac muscle is SERCA2a, and its alternative splicing product, SERCA2b, is expressed in smooth muscle, and thus in most all tissues at relatively low levels.15 17 18 SERCA2a is also expressed in relatively high levels in slow skeletal muscle but its density of expression is lower than SERCA1a in fast skeletal muscle.16 19 Known functional differences among the isoforms are that SERCA2b has lower intrinsic ATPase activity and Ca2+ uptake rate than the other isoforms20 and SERCA3, ubiquitously expressed in low levels,21 has a lower Ca2+ affinity than the other isoforms.20
The purpose of the present investigation was to examine whether the prolongation of SR Ca2+ cycling in skeletal muscle in CHF is associated with changes in the expression of SERCA. Northern and ribonuclease protection assays were used to measure expression of the SERCA1 and SERCA2 gene products in several different skeletal muscle types from rats with CHF. SERCA2 gene products were measured in cardiac muscle for comparison. Western blotting was used to examine SERCA1 and SERCA2 protein expression and SR Ca2+ ATPase activity was measured to determine functional enzyme activity. We found significant decreases in SERCA1a mRNA and protein expression as well as in SR Ca2+-ATPase activity. The data are consistent with the idea that a common factor (or factors) associated with the CHF condition may target striated muscle in general, acting as a regulator of SERCA expression and consequently altering function.
Materials and Methods
A well-characterized genetic rat model of CHF was used for these studies.22 23 Animals were bred for Spontaneous Hypertension and Heart Failure (SHHF/Mcc-facp). In brief, the rat colony originated from a cross between a Koletsky and an SHR from the SHR/N colony at the National Institutes of Health. Backcrosses (at least 14) were conducted at the NIH to produce a congenic SHR/N-cp strain with elimination of Sprague-Dawley genes. The Institute of Laboratory Animal Resources, National Research Council, subsequently (1990) replaced the term SHR with SHHR because of 11 years of selective breeding to favor the early and reproducible development of heart failure. The facp indicates that some members (25%) of the colony carry the “corpulent” gene. The reproducible development of dilated cardiomyopathy in the SHHF/Mcc-facp is unique among animal models of spontaneous hypertension, and it has been maintained for 28 generations. The genetics of CHF are multifactorial and the responsible genes have not yet been characterized. All SHHF rats are hypertensive and develop spontaneous CHF in the second year of life (males earlier than females, obese earlier than lean). Since the SHHF rat possesses both SHR and Sprague-Dawley genes, neither the Wistar-Kyoto nor the Sprague-Dawley is a perfect normal control. The Wistar-Furth is most commonly used as a control because of its many similarities to the SHHF.
Clinical signs of CHF include generalized and subcutaneous edema, hydrothorax, dyspnea, cyanosis, orthopnea, enlarged heart, left atrial thrombosis, and hyperemia of lungs, liver, and kidney. The left ventricle hypertrophies early in life in response to hypertension, then dilates in response to CHF, as does the right ventricle. Biochemical changes include reversion of heart contractile proteins to fetal isoforms, V1 to V3 shift in heart native myosin expression (during hypertrophy and at failure), elevated plasma and heart tissue atrial natriuretic factor, and increased plasma renin and norepinephrine in both lean and obese rats. The drugs captopril, enalapril, nifedipine, and felodipine all delay heart enlargement and myosin shift, whereas verapamil triggers overt failure. ACE inhibitors decrease circulating angiotensin II, and the initial decrease in blood pressure appears to be related to circulating plasma renin activity. A progressive functional decompensation has been noted in ECG patterns, in hemodynamic studies (depressed +dP/dT and increased end-diastolic pressure in both ventricles), and by echocardiographic analysis. Postmortem examination shows dilation of all cardiac chambers, thickened right and left ventricular free walls, hepatomegaly with ascites, and pulmonary edema with pleural effusion. The progression of the disease as well as the pathological alterations in SHHF mimics many of the findings in human CHF patients, particularly human hypertensive subjects who eventually develop CHF.
Five different sets of control and CHF rats (groups A, B, C, D, and E) were used for the present study (Table 1⇓). More than one set of rats was used because not all tissues were available from all sets of rats and not all measurements could be made on any one set of rats. Also, since true (same strain) age-matched controls are not possible using SHHF rats, we used several different types of control rats to ensure that changes in variables studied were due to the CHF condition rather than to age or strain differences of the controls. The female SHHF rats in failure were typically 3 months older than male SHHF rats in failure. When SHHF rats were used as controls (groups A, B, and some in C), they were necessarily younger than the SHHF rats in failure due to the progression of failure with age. Different strains of rats were used as control in groups D, E, and some in C to more closely age-match the SHHF rats in failure. However, the type of control did not influence the changes observed in failure. Several of the assays for gastrocnemius, diaphragm, and TA muscles were performed on more than one set of rats. A summary of all the assays done on all muscles by group is given in Table 2⇓.
SHHF rats were identified as being in end-stage failure when they displayed dyspnea, piloerection, cold tail, akinesis, and anorexia. At the time of failure, each rat was taken and anesthetized along with a control rat for dissection of tissues (see below). The clinical signs of the rats in failure, as detailed above, were noted during dissection.
To obtain muscles, rats were anesthetized with sodium pentobarbital administered intraperitoneally (40 mg/kg). Rats in CHF received about half the normal dosage because they reached deep anesthesia with less drug. Muscles were removed and immediately weighed and either frozen in liquid nitrogen or homogenized in buffers as indicated below. Rats were euthanized by removal of the heart during deep anesthesia.
Crude membrane, SR vesicles, or postnuclear homogenates were isolated from whole muscle as we have previously described.10 24 Recovery of SERCA from whole muscle in the postnuclear homogenates is 95% to 99%.16 Protein concentration was determined in SR vesicles and crude membranes with a kit (BioRad) based on the Bradford method. In postnuclear homogenates, protein concentration was determined using a detergent comparable assay (BioRad).
Immunoblotting was carried out as we have detailed previously,24 with A52, a monoclonal antibody specific for SERCA1 (a gift from Dr D. MacLennan25 ), 7E6, a monoclonal antibody specific for SERCA2 (a gift from Dr L. Jones26 ), or C4, a polyclonal antibody that recognizes all SERCA isoforms (a gift from Dr J. Lytton20 ). However, in the present study a different secondary antibody and detection system were used. Goat anti-rabbit IgG or goat anti-mouse IgG conjugated to horseradish peroxidase were used as secondary antibodies. A chemiluminescent substrate-detection system (KPL) was used to produce a light signal that was recorded on x-ray film. Signals on film were quantified by laser densitometry.
SR Ca2+-stimulated ATPase activity was measured in isolated membranes using an enzyme-coupled assay as detailed previously.10 Ca2+-stimulated ATPase activity was measured as the difference between total ATPase activity and basal (Mg2+-dependent) ATPase activity. Mg2+-dependent ATPase activity was not different between control and CHF groups (not shown), indicating that the calculated differences were due to Ca2+-stimulated rather than Mg2+-dependent ATPase activity.
DNA and RNA Probes
BstXI digestion (at 3244-3455 bp) of RtSkm3,16 a rat SERCA1a cDNA, yielded a 211 bp fragment from the 3′UTR, which was used to detect both SERCA1a and SERCA1b mRNA. This fragment does not distinguish the two isoforms on Northern blots because the transcript size differs by only 42 bp. To distinguish SERCA1a from 1b, a probe was made for use in a ribonuclease protection assay. The plasmid containing the cDNA was linearized with BglII (at 3081 bp) and in vitro transcribed to yield a 424 nt transcript containing all of the 3′UTR. This probe protects a 374 nt fragment and a 270 nt fragment, corresponding to SERCA1a and SERCA1b, respectively, when incubated with skeletal muscle RNA and treated with a ribonuclease.
For Northern blots of SERCA2a, a Nsi I-Cla I digestion (at 3495-3864 bp) of a rat SERCA2a cDNA, clone RS8-17,27 yielded a 368 bp fragment corresponding to the 3′UTR, which recognizes SERCA2a and two processing intermediates of SERCA2b mRNA, which can all be distinguished by size on Northern blots.21 For concomitant quantification of mature SERCA2a and SERCA2b in a protection assay, a 258 bp fragment (at 2341-2599 bp) of clone RS8-17 (in pBR322) was amplified by PCR and subcloned into pBluescript. Linearization of this subclone with HindIII and in vitro transcription yielded a 359 nt transcript. In the protection assay, this probe protected a 258 nt fragment corresponding to SERCA2a, a 107 nt fragment corresponding to SERCA2b (including the two processing intermediates), and a 151 nt fragment corresponding to only the processing intermediates of SERCA2b but not mature SERCA2b.
The α1-subunit of the DHP receptor mRNA was detected on Northern blots using a 1000 bp EcoRI digestion fragment from the 3′ end of the rat cDNA clone, 1Fb.28 A 900 bp PstI-KpnI digestion fragment consisting of the coding region of a chicken β-actin cDNA29 was used for comparison to other mRNA signals on some Northern blots. This fragment cross-reacts with sarcomeric α-actin. In some cases, an oligonucleotide (30-mer) complementary to rat 18S ribosomal RNA was used as a comparison for mRNA signals on Northern blots. The 18S oligo was 5′ end-labeled using a kinasing reaction with T7 polynucleotide kinase and [γ-32P]dATP under conditions indicated by the manufacturer (New England Biolabs).
DNA probes for Northern analysis were labeled with the use of random priming with [α-32P]dCTP according to instructions of the manufacturer (NEB). For protection assays, antisense RNA probes were transcribed in vitro from linearized plasmid DNA templates with T3 and T7 bacteriophage RNA polymerases, according to instructions of the manufacturer (Ambion). Transcripts were labeled with 32P-labeled UTP. Full-length transcripts were isolated from prematurely terminated products and unincorporated nucleotides by gel purification.
Total cellular RNA was isolated from muscles using the single-step guanadine method.30 Particular care was taken to use acid phenol to obtain DNA-free RNA. Standard protocol was used for Northern blot analysis,31 as we have previously described.10
For ribonuclease protection assays, gel-purified antisense RNA probes were incubated with sample RNA under conditions that favor hybridization of complementary transcripts, essentially as described by the manufacturer (Ambion). After hybridization, the mixture was treated with T1 ribonuclease to degrade single-stranded, unhybridized probe. Labeled probe hybridizes to complementary RNA such that double-stranded hybrids are protected from ribonuclease digestion. Products were then separated on a polyacrylamide gel and visualized by autoradiography.
Signals on x-ray film from Northern blots and protection assays were quantified by laser densitometry. Plasmid amplification, purification, and subcloning were performed according to standard protocols.31
An unpaired Student’s t test was used to determine statistical significance, with P<.05 or P<.01 as indicated.
Skeletal muscle mass was assessed only from rats in groups D and E. This is because tissue from groups A, B, and C were shipped frozen such that accurate wet mass could not be determined. The mass of the right plus left ventricle of CHF rats in all groups was double that of controls (Table 1⇑). In skeletal muscles that express predominantly (≈80% to 95%) fast myosin and “fast” SR Ca2+-ATPase (SERCA1a), there was mild atrophy. Specifically, the plantaris, TA, and medial and lateral heads of the gastrocnemius were 16%, 13%, 14%, and 18% lower in CHF rats (P<.05). Conversely, in the diaphragm, a muscle that expresses more (≈20% to 80%) slow myosin, “slow” SR Ca2+-ATPase (SERCA2a), and is chronically active, there was no significant change in muscle mass. There was no statistical difference in body mass of control versus CHF rats. Therefore, muscle mass differences were maintained when normalized to body mass.
SR Ca2+-ATPase Activity
There was a 38%, 40%, and 38% decrease in SR Ca2+-ATPase activity with CHF in TA SR vesicles, gastrocnemius crude membrane, and diaphragm crude membrane, respectively (Fig 1⇓). The decreased ATPase activity was not seen in the SHHF rats until failure but was otherwise independent of the age or strain of the controls. Because there were no sex-related differences, values for males and females were analyzed together.
All SR and crude membrane samples used for ATPase activity and Western analysis were visualized on a 5% to 18% gradient SDS-polyacrylamide gel stained with Coomassie blue. Visually, all samples had very little or no contaminants such as myosin and actin, and there was no qualitative difference in banding pattern between control and CHF samples. Quantitatively, the stained band that putatively represents SERCA appeared lighter in the CHF versus the control lanes. Western blot analysis confirmed this observation as detailed below. There was no difference in the yield of SR or crude membranes from control versus CHF muscles (eg, average value for crude membrane, 3.5 mg/g wet muscle mass).
Western blot analysis of the same SR vesicles or crude membranes used for ATPase activity showed a 46%, 42%, and 36% decrease in the expression level of SERCA1 protein for TA, gastrocnemius (Fig 2⇓), and diaphragm muscles (Table 2⇑). Western analysis of a second set of diaphragm muscles from which postnuclear protein was isolated showed a 64% decrease in SERCA1 expression (Fig 2⇓). The difference in the magnitude of decrease in SERCA1 expression in postnuclear (64%) versus crude membrane (36%) samples is likely a result of muscle samples from different rats being used. Other group-specific differences in the magnitude of decrease include SERCA1 mRNA expression of the TA and diaphragm in group B (45% and 58% decrease, respectively) versus group E or D (26% and 37% decrease, respectively) (Table 2⇑).
Though we used a polyclonal antibody (C4) that recognizes all SERCA isoforms for the Western blots shown in Fig 2⇑, the decreases represent SERCA1 for the following reasons. In TA and gastrocnemius muscles, SERCA2 levels were below the detection limit of the sensitive isoform-specific monoclonal antibody 7E6. In the case of the diaphragm there was no difference in SERCA2 protein expression in CHF, determined with the use of 7E6 (Table 2⇑), so the differences are attributed to SERCA1. In addition, in a different group of diaphragm samples (group A) the 36% decrease in SERCA1 protein expression was observed on a Western blot with the use of A52, the SERCA1-specific antibody (Table 2⇑).
To establish that SERCA2a protein expression is decreased in the hypertrophied and failing heart of the SHHF rat as it is in other etiologies of CHF,13 a Western blot of postnuclear samples from ventricular muscle was performed. SERCA2 expression was 36% lower in CHF samples (Fig 3⇓).
Using Northern analysis, we showed a 45% and 43% decrease in SERCA1 expression in the TA and plantaris muscle with CHF, respectively (Fig 4⇓). However, gastrocnemius SERCA1 was unchanged compared with controls in two different groups analyzed (Fig 4⇓). Diaphragm muscle showed a 58% decrease in SERCA1 expression (Fig 4⇓). There was no change in SERCA2a (cardiac isoform), dihydropyridine (DHP) receptor (α1-subunit), or α-actin mRNA expression in diaphragm muscle with CHF (Fig 5⇓). Protection assays of RNA from diaphragm (Fig 6⇓) and TA muscles (Table 2⇑) showed that the decrease in SERCA1 was due almost entirely to a decrease in SERCA1a. In the diaphragm muscle, there was a 160% increase in the alternatively spliced neonatal SERCA1b isoform that cannot be seen at the exposure shown. However, the relative expression of SERCA1b is very low in control adult skeletal muscle,16 making even this large percent increase functionally insignificant. Protection assay of TA and diaphragm RNA also showed no difference in SERCA2a or SERCA2b, the alternatively spliced SERCA2 isoform (Table 2⇑). These data indicate that the CHF-induced change was SERCA1 isoform specific.
Ribonuclease protection assay showed that SERCA2a mRNA expression was decreased by 61% in cardiac ventricles of CHF rats compared with controls, with a 27% decrease in the expression of the alternatively spliced transcript, SERCA2b (Fig 7⇓). However, SERCA2b represents only ≈5% of the total SERCA expressed in cardiac muscle.16 Northern blot analysis of SERCA2a expression normalized to 18S rRNA showed a similar decrease in ventricular RNA samples from CHF rats (Table 2⇑). The decreased expression of SERCA2 protein and mRNA in the SHHF heart are consistent with its mechanical deficits.32 33 As with the protein expression data, these results are consistent with that seen in other etiologies of CHF (reviewed in Reference 1313 ).
A major component of myocardial dysfunction in CHF is the prolongation of intracellular Ca2+ transients associated with excitation-contraction coupling. This defect is thought to be due in part to decreased expression of SERCA, the rate-limiting protein of intracellular Ca2+ sequestration.13 Consequently, the failing heart has slowed SR Ca2+ uptake, Ca2+-ATPase activity, and relaxation (ie, diastole) characteristics.13 An intriguing observation is that similar to cardiac muscle, skeletal muscle in rats with infarction-induced CHF also have prolonged intracellular Ca2+ transients during twitch and tetanic contractions, accelerated fatigue development,7 slowed rates of tension developmentand relaxation,8 and decreases in SR Ca2+ uptake and ATPase activity.9 Data reported in the present study indicate that these skeletal muscle defects are at least in part due to decreases in the protein expression of the predominant skeletal muscle SERCA isoform. SR Ca2+-ATPase activity, a functional correlate of SERCA expression, was also significantly decreased in all muscles studied.
We studied several different skeletal muscle types to examine the generality of defects in SERCA expression because of differences in chronic recruitment patterns and in SERCA composition. Data from the present study and from the literature show that adult diaphragm muscle expresses ≈75% SERCA1a, 23% SERCA2a, and low levels of SERCA1b, SERCA2b, and SERCA3, whereas adult TA muscles express ≈95% SERCA1a, 3% SERCA2a, and the balance SERCA2b, SERCA1b, and SERCA3.16 21 Gastrocnemius and plantaris muscles have similar SERCA isoform distribution to the TA but with slightly greater SERCA2a expression.24 31 In the plantaris, TA, and diaphragm muscle, there was a significant decrease in SERCA1a mRNA. SERCA1 protein expression and SR Ca2+-ATPase activity, assayed in TA and diaphragm muscle, were also decreased. These data suggest pretranslational control of gene expression, as observed with the SERCA2 gene expression in the failing heart. In gastrocnemius muscle, SERCA1 mRNA was unchanged, whereas SERCA1 protein expression and SR Ca2+-ATPase activity decreased. These observations suggest translational control of SERCA1 expression. CHF did not induce alterations in the expression of the SERCA2 gene products or of a major component of the Ca2+ release mechanism, the α1-subunit of the DHP receptor.
The mechanism by which CHF alters muscle gene expression is unknown, but physical inactivity secondary to CHF has been suggested as a cause of skeletal muscle abnormalities.4 However, recent evidence does not support this idea.6 With respect to the decreases in SERCA1 expression and the related functional changes, inactivity is unlikely to be a contributing factor because it leads to changes opposite to those found in heart failure. That is, muscular inactivity due to the removal of weight-bearing leads to marked increases in SERCA1 expression at the mRNA and protein level and increases in SR Ca2+-ATPase activity, SR Ca2+ uptake rate,34 and rates of muscle relaxation10 11 ; these are opposite to the observations in the present study with CHF. Finally, the fact that deficits in SERCA expression shown herein and the related mechanical properties8 are manifest in the diaphragm, a muscle that is continuously active to support breathing, also suggests that the deficits are independent of any physical inactivity associated with CHF.
Although the trigger of functional defects in skeletal and cardiac muscle in CHF remains unknown, decreased SERCA2a expression in the hypertrophied heart may be a phenotypic adaptation to overload similar to the decrease in SERCA1 expression in work-induced hypertrophy of skeletal muscle.24 However, the scenario is more complex when the hypertrophied heart goes into failure. At this point, the prolonged intracellular Ca2+ movements and decreased SERCA2a expression are exacerbated.35 A major finding of the present study is that significant decreases in SERCA1a expression occur in skeletal muscle where there is no hypertrophy. Also, the decrease in SERCA1a expression in skeletal muscle does not occur until frank failure, corresponding with the exacerbation of decreased SERCA2a expression in the heart. Together, these data suggest that aspects of the CHF condition other than hypertrophy affect SERCA expression and Ca2+ cycling defects in skeletal and possibly cardiac muscle.
Since SR Ca2+ cycling defects are similar in cardiac and skeletal muscle in CHF, it has been suggested that a common factor may be involved in triggering them.7 Results in the present study further support this idea and suggest that such a factor may be an upstream regulator of SERCA gene expression. There are elevations in a number of circulating hormones/factors with most etiologies of CHF; those characterized in the SHHF rat with CHF are norepinephrine, tumor necrosis factor, atrial natriuretic peptide, and renin-angiotensin.22 23 Activation of the local renin-angiotensin system (RAS) appears to be involved in the regulation of growth and some phenotypic changes in cultured hypertrophying cardiomyocytes and in the hypertrophied heart.36 37 38 Some evidence suggests that activation of the systemic RAS correlates with the transition from the hypertrophied to failing heart.39 This is consistent with our observations of skeletal muscle abnormalities manifest in failure. Another study showed that elevated norepinephrine levels in CHF downregulate β1-adrenergic receptors in the heart and in vascular smooth muscle at the mRNA and protein level.40 41 This effect appears to be mediated by decreases in the stability of the β1-adrenergic receptor mRNA by induction of AU-rich, mRNA-binding proteins.40 Similarly, elevated norepinephrine could also be an upstream regulator affecting SERCA expression.
In summary, results of this study showed that similar to the failing heart, the abnormalities related to excitation-contraction coupling in skeletal muscle with CHF are associated with decreases in SERCA expression. In most cases this was due to decreases at both the protein and mRNA level of the predominant skeletal muscle isoform and decreases in SR Ca2+-ATPase activity. These changes appear to be a primary change associated with the CHF condition unrelated to physical activity level. A common circulating factor may be involved in regulating decreased SERCA expression in skeletal as well as cardiac muscle in CHF.
Selected Abbreviations and Acronyms
|CHF||=||congestive heart failure|
|SERCA||=||sarcoplasmic (or endoplasmic) reticulum calcium ATPase|
|SHR||=||spontaneously hypertensive rat(s)|
|SHHF||=||spontaneous hypertension and heart failure|
This study was supported by a Grant-in-Aid and an Established Investigator Award from the American Heart Association (Dr Kandarian) and by US Public Health Service Grants AR-41705 (Dr Kandarian) and HL-48835 (Dr McCune). Salaries and research support was provided in part by state and federal funds appropriated to the Ohio Agricultural Research and Development Center, Ohio State University (Dr McCune).
- Received April 30, 1997.
- Accepted August 26, 1997.
- © 1997 American Heart Association, Inc.
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