Effect of heart failure on skeletal muscle myofibrillar protein content, isoform expression and calcium sensitivity

International Journal of Cardiology 107 (2006) 211 – 219 www.elsevier.com/locate/ijcard

Effect of heart failure on skeletal muscle myofibrillar protein content, isoform expression and calcium sensitivity Michael J. Toth*, Bradley M. Palmer, Martin M. LeWinter Departments of Medicine and Molecular Physiology and Biophysics, University of Vermont, Burlington, VT 05405, United States Received 10 December 2004; received in revised form 28 January 2005; accepted 11 March 2005 Available online 28 April 2005

Abstract Background: Alterations in skeletal muscle with heart failure contribute to exercise intolerance and physical disability. The majority of studies to date have examined abnormalities in skeletal muscle oxidative capacity and mitochondrial function. In contrast, less information is available regarding the effect of heart failure on myofibrillar protein metabolism and function. To address this issue, we examined the effect of heart failure on skeletal muscle myofibrillar protein content, isoform distribution and Ca2+ sensitivity. Methods: We measured skeletal muscle myosin heavy chain (MHC) and actin protein content and MHC isoform distribution in soleus (SOL), extensor digitorum longus (EDL), plantaris (PL) and diaphragm (DIA) muscles and myofibrillar Ca2+ sensitivity in EDL muscles from Dahl salt-sensitive rats with (high-salt fed: HS; n=10) or without heart failure (low-salt fed: LS; n=8) and assessed the relationship of these variables to markers of disease severity. Results: No differences in muscle mass were found. Similarly, no differences in MHC (mean T SE; SOL: 1353 T 29 vs. 1247 T 52; EDL: 1471 T 31 vs. 1441 T 31; PL: 1207 T 66 vs. 1286 T 36; DIA: 1166 T 42 vs. 1239 T 26 AU/Ag protein) or actin (EDL: 348 T 13 vs. 358 T 19; PL: 245 T 20 vs. 242 T 9; DIA: 383 T 9 vs. 376 T 17 AU/Ag protein) protein content or the actin-to-MHC ratio were observed, with the exception of lower ( P<0.01) actin content in the soleus of LS rats (352 T 7 vs. 310 T 8 AU/Ag protein). MHC isoform expression (I, IIa, IIx, IIb) did not differ between groups in SOL (I: 89 T 1% vs. 85 T 2%; IIa: 11 T1% vs. 15 T 2%), EDL (IIx: 43 T 10% vs. 38 T 10%; IIb: 57 T 10% vs. 62 T 10%), PL (I: 6 T 4% vs. 3 T 3%; IIa: 1 T1% vs. 1 T1%; IIx: 31 T 3% vs. 26 T 4%; IIb: 62 T 5% vs. 71 T 6%) or DIA (I: 43 T 6% vs. 36 T 6 %; IIa: 9 T 1% vs. 7 T 1%; IIx: 47 T 6% vs. 56 T 7%; IIb: 2 T 1% vs. 1 T 0.5%) muscles. Moreover, heart failure did not affect the Ca2+ sensitivity (i.e., pCa50) of extensor digitorum longus myofilaments (5.68 T 0.11 vs. 5.65 T 0.09). Finally, MHC and actin content, MHC isoform distribution and myofibrillar Ca2+ sensitivity were not related to markers of disease severity. Conclusions: Our results show that this animal model of heart failure is not characterized by alterations in the quantity or isoform distribution of key skeletal muscle myofibrillar proteins or the Ca2+ sensitivity of isometric force production. These findings suggest that alterations in skeletal muscle myofibrillar protein metabolism do not develop in parallel with myocardial failure in the Dahl saltsensitive rat. D 2005 Elsevier Ireland Ltd. All rights reserved. Keywords: Myosin heavy chain; Actin; Isoform; Exercise intolerance

1. Introduction Skeletal muscle adaptations play an important role in the symptomology of chronic heart failure [1]. The * Corresponding author. Health Science Research Facility 126B, University of Vermont, Burlington, VT 05405, United States. Tel.: +1 802 656 7989; fax: +1 802 656 0747. E-mail address: [email protected] (M.J. Toth). 0167-5273/$ - see front matter D 2005 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.ijcard.2005.03.024

majority of studies that have examined alterations in skeletal muscle have focused on changes in oxidative capacity and mitochondrial function [2– 4]. In contrast, relatively less is known about the impact of heart failure on myofibrillar protein metabolism. Several rodent models have been used to characterize the effect of heart failure on myofibrillar proteins. Most of these investigations have focused on changes in fiber type or myosin heavy chain (MHC) isoform expression.

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Results have been equivocal. Although many studies have observed alterations in fiber type or isoform distribution with heart failure [5 – 9], no consistent pattern has emerged. In fact, in some muscles, divergent results have been noted between studies [5 – 7,9]. Findings from Simonini et al. [7] have raised the intriguing possibility that heart failure might also affect the quantity of MHC in skeletal muscle, an alteration that would have clear functional significance. No subsequent studies, however, have measured skeletal muscle MHC protein content. Because of these inconsistencies and the paucity of information on certain aspects of myofibrillar protein, a primary goal of our study was to examine the effect of heart failure on skeletal muscle myofibrillar protein, with a specific emphasis on skeletal muscle MHC protein content and isoform distribution. Heart failure is characterized by skeletal muscle contractile dysfunction [1]. Several studies have identified defects in excitation – contraction coupling in skeletal muscle from rats with heart failure [10 – 12] which might contribute to contractile dysfunction. These alterations are similar to those observed in cardiac muscle [13,14], leading some investigators to hypothesize that heart failure is characterized by a generalized myopathy of striated muscle [10,11]. In addition to defects in excitation – contraction coupling, heart failure is characterized by changes in cardiac myofibrillar protein function; most notably, an increase in Ca2+ sensitivity [15,16]. Thus, if a generalized myopathy of striated muscle exists, skeletal muscle myofibrillar Ca2+ sensitivity should be increased similarly. Whether skeletal muscle myofibrillar Ca2+ sensitivity is affected by heart failure, however, is not clear. Thus, a secondary goal of our study was to examine the effect of heart failure on skeletal muscle myofibrillar Ca2+ sensitivity. The overall goal of our study was to evaluate the effect of heart failure on skeletal muscle myofibrillar protein metabolism and calcium sensitivity. To accomplish our objectives, we measured MHC and actin protein content, MHC isoform distribution and myofibrillar Ca2+ sensitivity in the Dahl salt-sensitive rat model of failure [17,18]. Myofibrillar protein content and isoform distribution measurements were performed on a variety of muscles (soleus, extensor digitorum longus, plantaris and diaphragm) that differ in both functional demands and isoform distribution. Myofibrillar Ca2+ sensitivity measurements were performed on the extensor digitorum longus muscle. This muscle was chosen because previous studies have shown that it exhibits altered excitation –contraction coupling similar to cardiac muscle [10,11]; whereas, other muscles, such as soleus, flexor digitorum brevis and gastrocnemius, do not [19,20]. An ancillary goal of our study was to examine whether adaptations in skeletal muscle myofibrillar protein metabolism with heart failure are related to markers of disease severity.

2. Materials and methods 2.1. Animals Dahl salt-sensitive rats were obtained (Taconic Inc.; Germantown, NY) at 6 weeks of age. All rats were in plastic-bottomed cages, two or three to a cage and were maintained on a 12-h:12-h light/dark cycle in a temperaturecontrolled room. Tap water and a low-salt (0.6%) chow were available ad libitum prior to and during the study. At 7 weeks of age, rats were divided into populations that received either high-salt (HS, 8% NaCl; n=10; 9 male, 1 female) or low-salt (LS, 0.6% NaCl; n=8; 4 male, 4 female) food pellets of similar macronutrient content. Six weeks following the initiation of diet, echocardiography was performed once per week in HS rats to assess left ventricular (LV) function. Myocardial failure was considered to be present if LV fractional shortening was  30% [16,18]. Skeletal muscle tissue was removed within 1 week of detection of heart failure. Echocardiography was performed on LS rats prior to collection of muscle tissue. All procedures were approved by the Institutional Animal Care and Use Committee of the University of Vermont and studies were conducted in accordance with the Guide for the Care and Use of Laboratory Animals. Data from a sub-set of these animals (n = 4 HS rats) have been published previously [21]. 2.2. Muscle tissue Rats were anesthetized using sodium pentobarbital (90 mg/kg, IP), a tracheotomy was performed and ventilation initiated with a respirator. Muscles were removed approximately 25 min following initiation of anesthesia. Soleus, extensor digitorum longus and plantaris muscles for myofibrillar protein content and MHC isoform distribution measurements were obtained from the left hindlimb, weighed and frozen in liquid N2. A sample of diaphragm muscle was removed and frozen immediately in liquid N2 for myofibrillar protein content and isoform distribution measurements. All four muscles were available from 8 HS and 7 LS rats for myofibrillar protein content and MHC isoform determinations. The extensor digitorum longus muscle from the right hindlimb was removed for functional measurements, as described below (HS: n = 6; LS: n = 7). Diaphragm muscle was carefully excised from a separate group of 15 rats (HS: n = 7; LS: n = 8) for muscle weight determinations. 2.3. Echocardiography Rats were anaesthetized with isoflurane (1%), placed in the supine position on top of a warming pad and their precordium shaved. Echocardiography was performed using a Sequoia system with a 15L8 linear array transducer (15.0-MHz; Acuson Corp., Mountain View, CA) [16].

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Briefly, in the two-dimensional mode, the left ventricle (LV) was imaged at the tip of the papillary muscle. This allowed positioning of the M-mode cursor perpendicular to the LV septum and posterior LV wall. M-mode images were recorded. LV dimensions (end-diastolic diameter: LVEDd, and end-systolic diameter: LVESd) were measured using the leading edge-to-leading edge convention. Posterior wall thickness was measured at end diastole. Values from all measured beats were averaged. Fractional shortening (%) was calculated as: ((LVEDd LVESd)/ LVEDd) * 100. 2.4. MHC and actin content MHC and actin content were determined by SDSPAGE, according to the method of Haddad et al. [22], with minor modifications. Muscle tissue was homogenized in extraction buffer (0.6 M KCl, 0.15 M KPi, 20 mM EDTA, 5 mM MgCl2, 3.3 mM ATP, pH 6.7) using a glass homogenizer and was incubated on ice for 90 min. The homogenate was centrifuged briefly (6500  g for 20 s) and analyzed for protein content using bovine serum albumin as a standard (Bio-Rad Laboratories, Hercules, CA). An aliquot of the muscle homogenate was added to loading buffer (2% SDS, 62.5 mM TRIS, 10% glycerol, 0.001% bromophenol blue, pH 6.8 with 5% h-mercaptoethanol) and heated for 5 min at 100 -C. For each muscle sample, 2 Ag of protein was loaded per lane, which was within the linear range of detection. A representative gel and data showing the linearity of these measurements through a range of protein loads is provided in Fig. 1. The stacking gel contained 4% acrylamide-N,N¶-methylene-bisacrylamide (bis) and the resolving gel 10% acrylamide-bis. Gels were run at constant current (60 mA) for approximately 2.5 h at 22 -C and then were stained with Coomassie blue. MHC and actin band intensity were determined by densitometry (Quantity One; Bio-Rad Laboratories; Hercules, CA) and were expressed as densitometric units per Ag of protein loaded. All samples for each muscle (n = 15 rats) were run on the same gel to eliminate gel-to-gel variability. All densitometric measurements were done by the same assessor who was blinded to treatment status (i.e., HS vs. LS). 2.5. MHC isoform distribution The relative distribution of MHC isoforms was determined according to the method of Klitgaard et al. [23] with minor modifications. Briefly, muscle tissue was homogenized in buffer (20 mM KCl, 2 mM K2HPO4, 1 mM EGTA, pH 6.8) and incubated on ice for 15 min. The homogenate was centrifuged at 12,000  g for 15 min, washed with homogenization buffer and centrifuged for 1 min at 12,000  g. The supernatant was decanted and the pellet extracted with 40 mM Na4P2O7, 1 mM MgCl2, 1 mM EGTA, pH 9.5 on ice for 15 min. The supernatant

Fig. 1. Linearity of MHC and actin protein content measurements by SDSPAGE. Panel A shows a representative gel with protein loads of 1, 2, 3 and 4 Ag. Panel B shows representative data for the linear response for MHC (h; r 2 = 0.995) and actin (g; r 2 = 0.997) content for these protein loads from n = 2 animals (1HS, 1LS).

was assayed for protein content using bovine serum albumin as a standard (Pierce Biotechnology; Rockford, IL), added to loading buffer and heated for 5 min at 100 -C. The myosin containing supernatants were analyzed by SDS-PAGE. The stacking gel contained 37.5% glycerol (w/v), 4% acrylamide-bis (50:1) and the resolving gel 37.5% glycerol and 6% acrylamide-bis (50:1). Gels were run at 70 V for 16 h followed by 4 h at 200 V for soleus and EDL muscles and at 275 V for 24 h for diaphragm and plantaris. Temperature throughout the entire run was maintained at 6 -C. Gels were stained with Coomassie blue (soleus and extensor digitorum longus) or silver stained (plantaris and diaphragm; SilverSnap II; Pierce Biotechnology; Rockford, IL) and the relative distribution of MHC isoforms quantified by densitometry (Quantity One; Bio-Rad Laboratories; Hercules, CA). All samples for each muscle (n = 15 rats) were run on the same gel to eliminate gel-to-gel variability. All densitometric measurements were done by the same assessor who was blinded to treatment status.

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2.6. Calcium sensitivity measurements

3.2. Body mass and skeletal muscle weights

Calcium sensitivity measurements were performed on chemically skinned extensor digitorum longus muscle. Prior to excision, the muscle was tied to a wooden stick, tendons were cut and the entire preparation placed in a skinning solution (1% Triton X-100 wt/vol and 50% glycerol wt/vol with 10 Ag/ml leupeptin) for 24 h at 4 -C. Following skinning, the muscle was removed and placed in storage solution (same as skinning solution minus Triton X-100) at 4 -C until analysis. Muscle fiber bundles (150 to 200 Am length) were dissected and mounted between a piezoelectric motor (Physik Instrument; Auburn, MA) and a strain gauge (Aurora Scientific; Richmond Hill, Ontario) at room temperature (22 -C) and stretched until passive tension was developed. This yields sarcomere lengths of approximately 2.5 Am in both HS and LS rats. We should note, however, that minor differences in starting sarcomere length that might arise using this approach would have minimal effects on pCa50 [24]. Solutions were calculated by solving equations for ionic equilibria [25] and concentrations are expressed in mM, unless otherwise specified: relaxing solution –pCa 9.0, 5.0 EGTA, 3.16 ATP, 1.0 Mg2+, 0.25 Pi, 12 phosphocreatine (PCr), 240 U/mL creatine kinase (CK), ionic strength 160, pH 7.1 and activation solution – same as relaxing solution with pCa 4.0. Prior to measurements, strips were maximally activated with pCa 4.5 and then relaxed. Strips were calcium activated at pCa 9 to pCa 4.5. Tension measures from multiple strips from each muscle were averaged to provide a single measure for that muscle. Data were fitted to the Hill equation using a nonlinear least squares algorithm.

Final body mass and skeletal muscle weights are shown in Table 1. No differences in body or muscle tissue masses were found.

2.7. Statistical analysis Differences between groups were determined by unpaired, Student’s t-test. Relationships between variables were determined by linear regression analysis. All data are expressed as mean T SE, unless otherwise specified.

3. Results 3.1. Indicators of cardiac failure LV fractional shortening was lower ( P < 0.01) in HS compared to LS rats (27 T 1% vs. 45 T 1%). Ventricular hypertrophy was evident as greater ( P < 0.01) total heart mass expressed per unit body mass in HS (n = 9) compared to LS rats (5.7 T 0.2 vs. 3.4 T 0.1 mg/g). Increased wet weight of the lung (7.3 T 0.6 vs. 4.4 T 0.2 mg/g; P < 0.01) and liver (44.0 T 1.0 vs. 24.4 T 1.2 mg/g; P < 0.01) in HS compared to LS (n = 7) rats expressed per unit body mass provides further evidence of decompensated failure. The average time on diet prior to study did not differ between groups (HS: 10.4 T 0.4 vs. LS: 9.5 T 0.4 weeks).

3.3. Myofibrillar protein content MHC and actin protein content are shown in Fig. 2. MHC content (AU/Ag protein) did not differ between HS and LS rats for soleus (HS: 1353 T 29 vs. LS: 1247 T 52), extensor digitorum longus (HS: 1471 T 31 vs. LS: 1441 T 31), plantaris (HS: 1207 T 66 vs. LS: 1286 T 36) or diaphragm (HS: 1166 T 42 vs. LS: 1239 T 26) muscles. Similarly, no differences in actin content (AU/Ag protein) were found in, extensor digitorum longus (HS: 348 T 13 vs. LS: 358 T 19), plantaris (HS: 245 T 20 vs. LS: 242 T 9) or diaphragm (HS: 383 T 9 vs. LS: 376 T 17) muscles. Actin content of the soleus muscle was lower ( P < 0.01) in LS (310 T 8) compared to HS (352 T 7) rats. No differences in the actin-to-MHC ratio were found in soleus (HS: 26 T 1% vs. LS: 25 T 1%), extensor digitorum longus (HS: 24 T 1% vs. LS: 25 T 1%), plantaris (HS: 21 T 2% vs. LS: 19 T 1%) or diaphragm (HS: 33 T 1% vs. LS: 30 T 2%) muscles. Neither MHC nor actin protein content nor actin/MHC were correlated to fractional shortening or relative heart, liver or lung weights in HS rats. 3.4. MHC isoform expression MHC isoform distribution patterns are shown in Fig. 3. In soleus, only MHC I and IIa isoforms were found. No difference in either isoform was noted between groups (I: HS: 89 T 1% vs. LS: 85 T 2%; IIa: HS: 11 T1% vs. LS: 15 T 2%). In extensor digitorum longus, MHC IIx and IIb were the only isoforms detected. No differences in either isoform was found between groups (IIx: HS: 43 T 10% vs. LS: 38 T 10%; IIb: HS: 57 T 10% vs. LS: 62 T 10 %). All four MHC isoforms were detected in plantaris and diaphragm

Table 1 Final body mass and skeletal muscle weights of HS and LS rats Variable

HS

LS

n

Final body mass (g) Soleus (mg) (mg/g) Plantaris (mg) (mg/g) Extensor digitorum longus (mg) (mg/g) Diaphragm (mg) (mg/g)

379 T 17 179 T 6 0.48 T 0.01 421 T 21 1.11 T 0.04 177 T 8 0.47 T 0.01 92 T 7 0.22 T 0.01

383 T 36 159 T 12 0.42 T 0.01 430 T 32 1.13 T 0.02 169 T 15 0.44 T 0.02 118 T 14 0.23 T 0.03

8/7 8/7 8/7 8/7 8/7 8/7 8/7 7/8 7/8

Muscle weight data are shown as absolute values (mg) and relative to body mass (mg/g). Values are mean T S.E. Body mass for rats on which diaphragm muscle weights were recorded are: HS: 413 T 23 vs. LS: 529 T 27; P<0.02 (see Muscle tissue for details).

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to fractional shortening or relative heart, liver or lung weights in HS rats. 3.5. Myofibrillar Ca2+ sensitivity Myofibrillar Ca2+ sensitivity of extensor digitorum longus muscles are shown in Fig. 4. Cross-sectional area of muscle fiber bundles were similar between HS and LS rats (83 T 22 vs. 88 T 23 mm2). No difference in the pCa50 was found (HS: 5.68 T 0.11 vs. LS: 5.65 T 0.09). The Hill coefficient did not differ between groups (HS: 3.80 T 0.95 vs. LS: 2.16 T 0.28). Neither pCa50 nor Hill coefficient data

Fig. 2. MHC and actin protein content of skeletal muscles. Panel A shows a representative gels for soleus (Sol), extensor digitorum longus (EDL), plantaris (Plant) and diaphragm (Dia) muscles for animals from HS (H) and LS (L) groups. Panel B to E shows average MHC and actin protein content data for soleus (B), extensor digitorum longus (C), plantaris (D) and diaphragm (E) muscle in HS (h; n = 8) and LS (g; n = 7) rats expressed per unit of protein loaded. Values for MHC are shown on the left axis and actin on the right axis. Values are mean T S.E. *, P < 0.01.

muscles. No difference in any isoform was found between HS and LS rats in either the plantaris (I: HS: 6 T 4% vs. LS: 3 T 3 %; IIa: HS: 1 T1% vs. LS: 1 T1%; IIx: HS: 31 T 3% vs. LS: 26 T 4%; IIb: HS: 62 T 5% vs. LS: 71 T 6%) or diaphragm (I: HS: 43 T 6% vs. LS: 36 T 6%; IIa: HS: 9 T 1% vs. LS: 7 T 1%; IIx: HS: 47 T 6% vs. LS: 56 T 7%; IIb: HS: 2 T 1% vs. LS: 1 T 0.5%). MHC isoform distributions were not related

Fig. 3. MHC isoform distribution of soleus (A), extensor digitorum longus (B), plantaris (C) and diaphragm (D) muscle in HS (h; n = 7) and LS (g; n = 8) rats expressed as a percentage of total MHC protein content. Representative gels showing separation of MHC isoforms for HS and LS rats are shown for each muscle. Values are mean T S.E.

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Fig. 4. Myofibrillar Ca2+ sensitivity of chemically skinned, extensor digitorum longus muscle fibers from HS (h; n = 6) and LS (g; n = 7) rats. pCa50 values did not differ between HS and LS rats (5.68 T 0.11 vs. 5.65 T 0.09, respectively). Values are mean T S.E.

were related to fractional shortening or relative heart, liver or lung weights in HS rats.

4. Discussion The present study focused on characterizing the effect of heart failure on skeletal muscle myofibrillar protein metabolism and function. No effect of heart failure on skeletal muscle MHC or actin protein content or MHC isoform distribution was observed. Moreover, no difference in the Ca2+ sensitivity of isometric force production was noted. Finally, variation in myofibrillar protein metabolism and function in rats with failure was not related to disease severity markers. Collectively, these results suggest that, in this animal model, heart failure does not significantly alter the metabolism or Ca2+ sensitivity of myofibrillar proteins. No differences in MHC or actin protein content were found, with the exception of lower actin content in the soleus muscle of LS rats. Because MHC and actin comprise a large percentage of total myofibrillar protein [26], these results imply that heart failure does not alter the relative amount of myofibrillar to total muscle protein content (i.e., MHC and actin content are expressed per Ag of total protein). Although actin content of the soleus was 12% lower in LS rats, no differences in the actin-to-MHC ratio was found between groups because of a corresponding 8% reduction in MHC content in LS rats. Because there were no differences in the actin-to-MHC ratio in any of the muscles studied, our findings further suggest no effect of heart failure on the stoichiometry of actin to myosin. Our results differ from those of Simonini et al. [7], who found a 30% reduction in MHC content of the soleus muscle in rats with heart failure compared to controls. This reduction was observed despite no difference in soleus protein content (non-significant 15% lower protein content in rats with heart failure), suggesting a selective reduction in MHC per unit

muscle protein. The reason for these differing results may relate to the animal models used. In the present investigation, Dahl rats were studied within a week of detection of failure. In contrast, Simonini et al. used a coronary artery ligation model and studied animals 20 weeks after surgery. Their paradigm likely resulted in a more chronic exposure of skeletal muscle to the heart failure syndrome. Thus, differences between studies may relate to variation in the duration of failure. Similarly, we cannot discount the possibility that differences between the two models in disease severity or other aspects of the heart failure syndrome (e.g., immune activation) contributed to divergent findings, although we should point out that the Dahl rat model is characterized by neurohumoral changes similar to human heart failure [27]. Because the phenomenon of selective MHC depletion can be caused by muscle disuse [22], the findings of Simonini et al. [7] could be explained by reduced physical activity, rather than an effect of heart failure, per se. Although Simonini et al. found no effect of heart failure on physical activity in this model in a subsequent study [28], it is important to note that activity measurements were conducted 8 weeks following surgery. Whether physical activity is reduced by 20 weeks in the coronary artery ligation model is not known. Numerous studies have examined the effect of heart failure on skeletal muscle MHC isoform expression. In general, these studies show that heart failure induces a shift in MHC isoform distribution. In peripheral skeletal muscle, a shift from a slow- towards a more fast-twitch phenotype has been noted [5– 9]; whereas, in diaphragm muscle, the reciprocal occurs [29,30]. Although this isoform shift is thought to be a hallmark of the skeletal muscle myopathy of failure [1], considerable disagreement exists among reports from animal models. For example, some studies have observed this isoform shift in soleus muscle [5,7,8], while others have not [6,9]. Differences among studies are not explained by the experimental model used since changes in soleus MHC isoform distribution have been observed in coronary artery ligation models in some studies [7], but not all [6,9]. The duration and degree of failure also probably do not explain differences among studies since Delp et al. [9] and Simonini et al. [28] reported differing effects on soleus muscle MHC isoform distribution despite the fact that, in both studies, heart failure was initiated by coronary artery ligation, infarct size and degree of failure were comparable and measurements were performed at identical times following surgery (8 weeks). In the present study, we found no effect of heart failure on MHC isoform expression in any muscle examined. Our results agree with Brunotte et al. [31], who found no effect of failure on fiber type distributions in calf muscles of rats 6 weeks following coronary artery ligation. Although it might be argued that our model is too acute to observe an effect of failure on MHC isoform, others have noted a switch in isoform distribution prior to the development of overt failure [32,33], suggesting that the skeletal muscle myopathy

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develops in parallel with cardiac failure. Thus, we believe that there was sufficient time for MHC isoform shifts to occur. The primary conclusion that can be drawn from these studies is that the effect of heart failure on skeletal muscle MHC isoform expression is highly variable. That factors such as the experimental model, duration of study and degree of failure do not explain divergent results indicates that other, as yet unidentified, variables contribute to shifts in MHC isoform distribution. One factor that might induce MHC isoform shifts is angiotensin II. Dalla Libera et al. [34] recently showed that angiotensin II blockade in a rat model of right-sided heart failure corrected shifts in skeletal muscle MHC isoform expression. The potential involvement of angiotensin II in regulating MHC isoform expression may explain why we did not observe an isoform shift. Dahl salt-sensitive rats have a polymorphism in the renin gene [35], which contributes to low tissue and plasma renin activity [36]. If angiotensin II plays an important role in driving changes in MHC isoform expression with heart failure, the genetic defect in the Dahl salt-sensitive strain may have limited skeletal muscle MHC isoform shifts. We should note, however, that recent studies have shown up-regulation of components of the renin – angiotensin system, as well as cardiac angiotensin II levels, in Dahl rats at the time of compensated left ventricular hypertrophy and failure [37]. Thus, despite the polymorphism in the renin gene, the Dahl model is characterized by activation of the renin –angiotensin system. To what degree this occurs in skeletal muscle, however, is not certain. Although alterations in excitation –contraction coupling have been noted in extensor digitorum longus muscle by several groups [10 – 12], our results suggest that this muscle is not characterized by changes in myofibrillar Ca2+ responsiveness. This result in skeletal muscle is in contrast to the increase in Ca2+ sensitivity observed in cardiac myofibrils in the Dahl rat model of failure by our laboratory [16], as well as in other animal models [15]. Thus, changes in skeletal muscle myofibrillar Ca2+ sensitivity do not occur in parallel with cardiac myofibrillar adaptations. Increased cardiac Ca2+ sensitivity is thought to arise, in part, from increased adrenergic stimulation [15]. Because the Dahl rat model of failure is characterized by neurohumoral activation and increased norepinephrine levels [27], it is unlikely that the absence of differences in pCa50 is due to lack of adrenergic stimulus. Instead, the reason for the discrepancy between cardiac and skeletal muscle myofibrillar Ca2+ sensitivity may relate to the fact that skeletal muscle isoforms of several key myofibrillar proteins lack sites for posttranslational modification that may regulate Ca2+ sensitivity. For example, although phosphorylation of cardiac troponin I via adrenergic signaling is thought to increase myofibrillar Ca2+ sensitivity [15], the skeletal muscle isoform of troponin I lacks these regulatory sites [38]. Our results agree with those of De Sousa et al. [8], who found no difference in pCa50 in either the soleus or

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gastrocnemius muscles of rats with heart failure compared to controls. Collectively, these data suggest that heart failure is not characterized by a generalized myopathy among striated muscles with respect to the Ca2+ sensitivity of isometric force production. The variability in results among studies regarding the effects of heart failure on myofibrillar protein metabolism highlights a limitation to the use of animal models. Although these models faithfully reproduce the myocardial properties of heart failure, there may be substantial variation in the effects on skeletal muscle. Because of this, extrapolation of results from animal models to humans should be made with caution. Close attention should be paid to the model system that is used and how it is applicable to human heart failure. In the present report, rats were studied shortly after the development of heart failure. In this context, our data reflect the acute effects of the transition from compensated cardiac hypertrophy to heart failure on skeletal muscle myofibrillar protein metabolism and function. This model allows us to examine the skeletal muscle adaptations that occur during the development of cardiac failure. We interpret our results to suggest that alterations in skeletal muscle myofibrillar protein do not develop in parallel with myocardial failure. Instead, skeletal muscle myofibrillar adaptations, such as decreased MHC protein content [7] and MHC isoform shifts [39,40], are likely related to prolonged exposure to aspects of the heart failure syndrome (e.g., neurohumoral activation). An alternative, and equally tenable, interpretation of our results is that they reflect the effect of heart failure on myofibrillar protein metabolism and function, uncomplicated by the secondary effects of physical inactivity and malnutrition-induced muscle atrophy. For example, in humans [41] and animal models, reduced physical activity may develop with chronic heart failure. Changes in myofibrillar proteins, therefore, may represent the combined effects of heart failure and muscle disuse. Because of this, our results may be a clearer representation of the primary effect of heart failure on skeletal muscle myofibrillar protein metabolism and function. We should also point out that our model has the advantage of not requiring a surgical intervention to induce failure. In such models, skeletal muscle adaptations may arise from muscle disuse during the post-surgical recovery period. Although appropriate shams ensure that all rats will undergo similar muscle changes, in those rats undergoing experimental manipulations, such as coronary artery ligation, circulatory and hormonal changes resulting from this intervention could function to maintain disuse-related skeletal muscle adaptations that develop in the post-surgical period. In this scenario, the myopathy is caused by muscle disuse and is preserved, rather than being promoted, by the development of failure. Our results suggest that, in the Dahl salt-sensitive model, heart failure is not characterized by alterations in skeletal muscle myofibrillar protein content, isoform distribution or Ca2+ sensitivity of isometric tension. These results suggest

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that, in its early stages, heart failure does not affect skeletal muscle myofibrillar protein content, isoform distribution or Ca2+ sensitivity.

Acknowledgments

[14]

[15] [16]

The authors would like to thank Jessica Saunders, Steve Bell, Ian Galbraith and Teruo Noguchi for their technical assistance. This work was supported by grants from the NIH (AM02125; AG-17494) and American Federation for Aging Research.

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