AJP - Heart Calcium Transients and Cell-Sarcomere
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Am J Physiol Heart Circ Physiol 293: H819-H828, 2007. First published April 20, 2007; doi:10.1152/ajpheart.00085.2007
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Diaphragm single-fiber weakness and loss of myosin in congestive heart failure rats

Hieronymus W. H. van Hees,1,5 Henricus F. M. van der Heijden,1,5 Coen A. C. Ottenheijm,1,5,6 Leo M. A. Heunks,1,2,5 Cindy J. C. Pigmans,1 Freek W. A. Verheugt,3 Rene M. H. J. Brouwer,4 and P. N. Richard Dekhuijzen1,5

Deptartments of 1Pulmonary Diseases, 2Intensive Care Medicine, 3Cardiology and 4Cardio-Thoracic Surgery and 5Institute for Fundamental and Clinical Human Movement Sciences, Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands; and 6Deptartment of Veterinary and Comparative Anatomy, Pharmacology, and Physiology, Washington State University, Pullman, Washington

Submitted 19 January 2007 ; accepted in final form 14 April 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Diaphragm weakness commonly occurs in patients with congestive heart failure (CHF) and is an independent predictor of mortality. However, the pathophysiology of diaphragm weakness is poorly understood. We hypothesized that CHF induces diaphragm weakness at the single-fiber level by decreasing myosin content. In addition, we hypothesized that myofibrillar Ca2+ sensitivity is decreased and cross-bridge kinetics are slower in CHF diaphragm fibers. Finally, we hypothesized that loss of myosin in CHF diaphragm weakness is associated with increased proteolytic activities of caspase-3 and the proteasome. In skinned diaphragm single fibers of rats with CHF, induced by left coronary artery ligation, maximum force generation was reduced by ~35% (P < 0.01) compared with sham-operated animals for slow, 2a, and 2x fibers. In these CHF diaphragm fibers, myosin heavy chain content per half-sarcomere was concomitantly decreased (P < 0.01). Ca2+ sensitivity of force generation and the rate constant of tension redevelopment were significantly reduced in CHF diaphragm fibers compared with sham-operated animals for all fiber types. The cleavage activity of the proteolytic enzyme caspase-3 and the proteasome were ~30% (P < 0.05) and ~60% (P < 0.05) higher, respectively, in diaphragm homogenates from CHF rats than from sham-operated rats. The present study demonstrates diaphragm weakness at the single-fiber level in a myocardial infarct model of CHF. The reduced maximal force generation can be explained by a loss of myosin content in all fiber types and is associated with activation of caspase-3 and the proteasome. Furthermore, CHF decreases myofibrillar Ca2+ sensitivity and slows cross-bridge cycling kinetics in diaphragm fibers.

skinned muscle single fibers; cross-bridge cycling kinetics; Ca2+ sensitivity; proteasome; caspase-3

THE MOST IMPORTANT CLINICAL manifestations of congestive heart failure (CHF) include impaired exercise capacity and dyspnea. Yet the severity of these symptoms poorly correlates with measures of left-ventricular performance in CHF patients (21). Interestingly, the degree of exercise intolerance is significantly correlated with peripheral muscle mass and function (11, 38, 59). Likewise, the sensation of dyspnea is closely related to respiratory muscle dysfunction (32). Moreover, maximal inspiratory pressure is reduced in patients with mild-to-moderate CHF (28) and has been identified as an independent predictor of survival in these patients (35). Although peripheral skeletal muscle alterations in CHF have been investigated in detail (11, 38, 59), relatively little is known about the pathophysiology of diaphragm weakness in CHF.

In line with reduced maximal inspiratory pressure in CHF patients, several animal studies of CHF described decreased specific force generation of diaphragm muscle bundles in vitro (29, 49, 50). Recently, Coirault et al. (12) found reduced in vitro motility of isolated myofilaments from the diaphragm of CHF rats, suggesting slower cross-bridge cycling kinetics. In addition, a rightward shift of the force-frequency relationship of CHF diaphragm bundles could involve a decreased Ca2+ sensitivity of force generation (52). Contractile properties are known to differ among fiber types in the diaphragm muscle (24, 25, 47). Because CHF has been shown to induce a fiber-type shift in the diaphragm (16, 27), changes in contractile properties could be ascribed to this fiber-type shift. However, to improve understanding of the pathophysiology of diaphragm weakness in CHF, diaphragm muscle function should be evaluated independently of a fiber-type shift, which has not been done yet. Skinned diaphragm single fibers provide an excellent model for studying myofilament function in the CHF diaphragm for every fiber type separately. The first hypothesis of the present study is that CHF decreases maximum force generation, slows cross-bridge cycling kinetics, and reduces Ca2+ sensitivity in skinned diaphragm single fibers.

Force-generating capacity of skinned muscle fibers depends on contractile protein content per half-sarcomere of the muscle fiber (6, 24). CHF is associated with peripheral skeletal muscle wasting (19), and, more particularly, loss of myosin is related to decreased leg muscle strength in CHF patients (57). In patients with chronic obstructive pulmonary disease, in experimentally induced hypothyroidism, and in unilateral denervation, decreased force generation of diaphragm fibers was explained by reduced myosin content per half-sarcomere (22, 23, 40). Myosin content in CHF diaphragm fibers has not been studied. The second hypothesis of the present study is that impaired force generation of skinned fibers is associated with reduced myosin content per half-sarcomere in CHF diaphragm fibers.

Selective degradation of myosin through the ubiquitin-proteasome pathway seems to play an important role in the loss of muscle mass in various diseases (1, 10). Caspase-3 activation has been proposed to be an initial step in the cleavage of the myofilaments, yielding fragments that are subsequently degraded by the ubiquitin-proteasome pathway (18). Our group has recently shown (40) that loss of myosin is a key event in the pathophysiology of diaphragm muscle weakness in patients with chronic obstructive pulmonary disease. Loss of myosin is associated with activation of the ubiquitin-proteasome pathway and the proteolytic enzyme caspase-3 in the diaphragm of these patients (39). It is yet unknown if activation of the proteasome and caspase-3 also occurs in the CHF diaphragm. The third hypothesis of the present study is that CHF enhances the proteolytic activity of caspase-3 and the proteasome in the diaphragm.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Experimental model. Adult male Wistar rats (260–300 g) were anesthetized by inhalation of an isoflurane-oxygen mixture (2–5% isoflurane), intubated, and mechanically ventilated. Ligation of the left coronary artery or sham operation were executed as described previously (44). In short, a left thoracotomy between the fourth and fifth rib was performed, exposing the left-ventricular wall. The left coronary artery was ligated at its origin by a 5-0 silk suture. Subsequently, the thorax was closed in three layers with 3-0 vicryl sutures. Sham operations were performed identically, without ligating the coronary artery. After the operation and during the following 2 days, buprenorphine (20 µg/kg sc) was administered daily for postoperative analgesia. Fifty coronary artery ligations and ten sham operations were performed. Perioperative survival was 42% in the ligated group and 100% in the sham-operated group and was comparable with previous studies using this model of heart failure (31, 56). At 14 wk after ligation or sham operation, the rats were anesthetized with pentobarbital (70 mg /kg ip) and were mechanically ventilated. Aortic and left-ventricular pressures were measured by a micromanometer-tipped catheter (SPC 330; Millar Instruments, Houston, TX) inserted through the right carotid artery. At the end of the hemodynamic measurements, a combined thoracotomy/laparotomy was performed, and the diaphragm, lungs, and heart were quickly excised. Heart weight was determined, and the heart was fixated in formalin for >72 h. Subsequently, the left ventricle of each heart was cut into three transverse sections in parallel with the atrioventricular groove. Transverse sections (7 µm thick) were cut and stained with Masson's trichrome stain. The size of the infarct areas was determined by planimetry, as described previously (44). Only data from rats with infarcts >35% of the left ventricle (n = 14) were analyzed in this study. Lungs were cleaned of fat and weighed before and after desiccation (3 days at 40°C). The diaphragm was divided in two parts: the right hemidiaphragm was used for determination of single-fiber contractile properties, and the left hemidiaphragm was used for caspase-3 and proteasome analysis. From seven CHF and six sham-operated rats, a part of the right hemidiaphragm was cut and weighed before and after desiccation (3 days at 40°C) to measure water content in the diaphragm muscle. This study was approved by the Animal Ethics Committee at Radboud University (Nijmegen, The Netherlands).

Diaphragm muscle single-fiber contractile measurements. Single-fiber contractile measurements and experimental protocol were performed according to previously described methods (40), with minor modifications, as described below.

Composition of solutions for single-fiber measurements. Relaxing solution consisted of (in mM) 1.0 MgCl2, 4.0 Na2ATP, 5 EGTA, 10 imidazole, and 15 creatine phosphate with sufficient KCl to adjust the total ionic strength to 150 mM at pH 7.0. The negative logarithm of the free Ca2+ concentration (pCa) of the relaxing solution was 9.0, whereas in activating solutions the pCa ranged from 7.0 to 4.0 (with maximal activation at pCa 4.0). To achieve appropriate pCa in activating solutions, sufficient CaCl2 was added to solutions. The composition of the solution for maximal rigor activation was the same as that of the pCa 4.0 solution, except that Na2ATP was omitted.

Diaphragm muscle single-fiber preparation and experimental setup. After excision, the diaphragm and adherent lower ribs were immediately submersed in cooled oxygenated (95% O2-5% CO2) Krebs solution at pH ~ 7.4. This Krebs solution consisted (in mM) of 137 NaCl, 4 KCl, 2 CaCl2, 1 MgCl2, 1 KH2PO4, 24 NaHCO3, and 7 glucose, with 25 µM D-tubocurarine (Sigma Aldrich, Bornem, Belgium). From the central costal region of the right hemidiaphragm, a rectangular bundle was dissected parallel to the long axis of the muscle fibers for single-fiber contractile measurements. The muscle bundle was pinned to cork and stored at 4°C in a relaxing solution containing 50% glycerol (vol/vol). After 24 h, the muscle strip was stored at –20°C for later analysis.

Approximately 1 h before determination of single-fiber contractile properties, the muscle bundle was transferred to relaxing solution (5°C) containing 1% Triton X-100 to permeabilize lipid membranes. From the muscle bundle, ~2-mm segments of single fibers were isolated by using microforceps. Subsequently, the fiber ends were attached to aluminum-foil clips and mounted on the single-fiber apparatus. Fibers were mounted in a temperature-controlled (20°C) flow-through acrylic chamber (120-µl volume) with a glass coverslip bottom on the stage of an inverted microscope (model IX-70; Olympus, Amsterdam, The Netherlands). Two stainless-steel hooks were used to mount the fiber horizontally in the chamber. One end of the fiber was attached to a force transducer (model AE-801; SensoNor, Horten, Norway) with a resonance frequency of 10 kHz, whereas the other end was attached to a servomotor (model 308B; Aurora Scientific, Aurora, ON, Canada) with a step time of 250 µs. In relaxing solution, sarcomere length was set at 2.4 µm as the optimal length for force generation (9, 61) with the use of a calibrated eyepiece micrometer. During experiments, sarcomere length was stabilized with the Brenner cycling method (5) as modified by Sweeney et al. (54). MIDAC software (Radboud University) and a data-acquisition board were used to record signals. Muscle-fiber length (~1–1.5 mm) was measured by using a reticule in the microscope eyepiece [x10 Olympus Plan 10, 0.30 numerical aperture (NA)]. The x-y fiber diameter (width) was measured with a x40 objective (x40 Olympus Plan 40, 0.60 NA). The x40 objective also was used to measure the x-z fiber diameter (depth) by noting the displacement of the microscope's objective while focusing on the top and bottom surfaces of the fiber. Three width and depth measurements were made along the length of the fiber, and the average values were used to calculate the fiber cross-sectional area, assuming that the fiber was ellipsoid in shape. Similar incidences of fibers that appeared injured (i.e., sarcolemmal damage, loss of cross-striation, or other irregularities) during microscopic examination were found in both groups, and those fibers were excluded from the study.

Diaphragm muscle single-fiber contractile determinations. Maximum specific force was determined by dividing the isometric force generated at pCa 4.0 by cross-sectional area. Maximum force at pCa 4.0 was also divided by the estimated value of myosin heavy chain content per half-sarcomere (see Myosin heavy chain isoform composition and content per half-sarcomere determination) to determine the force per half-sarcomere myosin heavy chain content.

The rate constant for tension redevelopment (ktr) was measured as described by Brenner and Eisenberg (7) during activation at pCa 4.0. In short, fibers were rapidly released by ~15% and then, ~50 ms later, restretched to their original length. During the rapid release and restretch, cross-bridges detach and force drops to zero. The cross-bridges then reattach, and force redevelops. The ktr value was determined by using a computer algorithm for least-squares fit of a first-order exponential.

Fiber stiffness was determined during sinusoidal length oscillations of 0.4% at 1 kHz. Stiffness provides an estimation of the number of strongly attached cross bridges (20). Stiffness was measured during Ca2+ activation at pCa 4.0 in the presence and absence (rigor solution) of ATP. Assuming that, during rigor activation, all cross-bridges are in the strongly attached state, the ratio of stiffness during Ca2+ activation to that during rigor activation ({alpha}fs) provides an estimate of the fraction of cross-bridges in the strongly attached state.

To determine force-pCa relationship, the isometric force generated in response to incubation with incremental Ca2+ concentrations (pCa 9.0, 7.0, 6.3, 6.0, 5.8, 5.5, 5.2, 5.0, 4.7, 4.5, and 4.0) was recorded. GraphPad Prism (GraphPad Software, San Diego, CA) was used to calculate the Ca2+ concentration required for half-maximum activation (pCa50), as an index of Ca2+ sensitivity of force generation, and the Hill coefficient, as a measure of myofilament cooperativity.

Diaphragm muscle single-fiber experimental protocol. Maximal specific force was determined by perfusing the experimental chamber at, successively, pCa 9 and 4.0. During the plateau phase of maximal activation at pCa 4.0, ktr was determined. Then pCa 9.0 solution was perfused through the chamber to relax the fiber.

Subsequently, fibers were perfused with solutions containing incremental Ca2+ concentrations. Each time a plateau was reached, the next pCa solution was perfused through the experimental chamber. When maximum force was reached at pCa 4.0, ktr was determined, followed by a determination of fiber stiffness. Subsequently, the fiber was perfused with rigor solution of pCa 4.0 to determine stiffness during rigor activation. Finally, the fiber was perfused at pCa 9 to verify baseline force.

Structural and functional instability of skinned single muscle fibers, reflected by an increased heterogeneity of sarcomere spacing during activation and reduced force generation over time, is a well-recognized problem (4). To improve functional stability of muscle fibers, we applied Brenner cycling (5) throughout the experimental protocol, except during determination of ktr, stiffness, and rigor. Accordingly, no functional decline in maximal force generation and ktr over time was observed.

Myosin heavy chain isoform composition and content per half-sarcomere determination. Determination of myosin heavy chain isoform composition and content by SDS-PAGE was adapted from Geiger et al. (24). Single fibers were detached from the force transducer and servomotor and were placed in 25 µl of SDS sample buffer containing 62.5 mM Tris·HCl, 2% (wt/vol) SDS, 10% (vol/vol) glycerol, and 0.001% (wt/vol) bromophenol blue at a pH of 6.8. The samples were stored at –20°C until they were assayed. The samples were denaturated by boiling for 2 min. The stacking gel contained a 4% acrylamide concentration (pH 6.8), and the separating gel contained 7% acrylamide (pH 8.8) with 30% glycerol (vol/vol). Control samples of rat diaphragm bundles were run on the gels for comparison of migration patterns of the myosin heavy chain isoforms. Sample volumes of 8 µl were loaded per lane. The gels were silver stained according to the procedure described by Oakley et al. (37). A standard curve of known contents of purified rabbit myosin heavy chain (M-3889; Sigma, Zwijndrecht, The Netherlands) was run on every gel to determine myosin heavy chain content in rat diaphragm muscle fibers.

After silver staining, the gels were scanned with an imaging densitometer and optical densities of the electrophoretic bands were quantified with GeneTools software (Syngene, Cambridge, UK). Background staining was subtracted from the density of the electrophoretic bands to determine the brightness-area product for each diaphragm muscle fiber. The relationship between the brightness-area product and myosin heavy chain content was linear across a range from 0.01 to 0.25 µg. The myosin heavy chain content in the loaded 8 µl SDS buffer was determined from the standard curve. Subsequently, the total myosin heavy chain content of the fiber (in 25 µl SDS buffer) was determined. Myosin heavy chain concentration was determined by dividing total myosin heavy chain content by fiber volume. Myosin heavy chain content per half-sarcomere, at a sarcomere length of 2.4 µm, was calculated through dividing fiber myosin heavy chain content by the amount of half-sarcomeres (2 x length of fiber/2.4).

Measurement of caspase-3 activity. Caspase-3 activity was determined as described previously (39). Frozen diaphragm samples were pulverized and homogenized on ice in a buffer containing 100 mM HEPES (pH 7.5), 10% sucrose, 0.1% Nonidet P-40, 10 mM dithiothreitol, and protease inhibitor cocktail (Sigma). Homogenates were subjected to three cycles of freeze-thaw before centrifugation at 18,000 g for 30 min. The supernatant (92.5 µg protein) was added to a reaction buffer consisting of 100 mM HEPES (pH 7.5), 10% sucrose, and 10 mM dithiothreitol. The fluorogenic substrate N-acetyl-Asp-Glu-Val-Asp-7-amido-4-methylcoumarin was then added, and the reaction was performed at 30°C for 60 min. The caspase-3 activity was determined by measuring the generation of the fluorogenic cleavage product (methylcoumarylamide) at 360 nM excitation wavelength and 460 nM emission wavelength with a spectrophotometer. Amido-4-methylcoumarin standards were used to quantify activity levels. Results were expressed as picomoles per minute per milligram protein. Addition of N-acetyl-Asp-Glu-Val-Asp-CHO (a caspase-3 inhibitor) to the reaction resulted in inhibition of methylcoumarylamide production, indicating the specificity of the assay for measuring caspase-3 activity.

Isolation of 20S proteasomes and measurement of proteasome activity. The 20S proteasome isolation and proteolytic activities were determined as described previously (39). To isolate 20S proteasomes, diaphragm samples frozen in liquid nitrogen were homogenized in ice-cold buffer (pH 7.5) containing (in mM) 50 Tris·HCl, 5 MgCl2, 250 sucrose, 1 1.4-dithiothreitol, and 0.2 PMSF, with protease inhibitor cocktail (Sigma) by means of a Dounce homogenizer. Subsequently, the proteasomes were isolated from the homogenates by three sequential centrifugation steps; the first centrifugation was at 10,000 g for 20 min. The supernatant was centrifuged at 100,000 g for 1 h. The obtained supernatant was then centrifuged at 100,000 g for 5 h. The final pellet containing the 20S proteasomes was resuspended in buffer (pH 7.5) containing 50 mM Tris·HCl, 5 mM MgCl2, and 20% glycerol. The protein content of the proteasome preparation was determined on the basis of a Bio-Rad protein assay (Bio-Rad, Veenendaal, The Netherlands). The proteolytic activity of the 20S proteasomes was determined by measuring the activity against the fluorogenic substrates succinyl-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin (LLVY) and N-carbenzoxy-Leu-Leu-Glu-7-amido-4-methylcoumarin (LLE) (Sigma). These substrates are preferentially hydrolyzed by the chymotrypsin-like and peptidylglutamyl peptidase activities of the 20S proteasome, respectively (15). To measure proteolytic activity, 15 µg proteasome extract was added to 60 µl medium containing 62.5 mM Tris·HCl, 12.5 mM MgCl2, 1.25 mM 1.4-dithiothreitol, 0.01 units apyrase, and 100 µM LLVY or 375 µM LLE. The reaction took place at 37°C for 45 min. The peptidase activity was determined by measuring the generation of the fluorogenic cleavage product (amido-4-methylcoumarylamide) at 380 nM excitation wavelength and 440 nM emission wavelength with a spectrophotometer. Standard curves were established for the fluorogenic product, and peptidase activity was expressed as picomoles per microgram protein per minute. Addition of MG-132 (a specific proteasome inhibitor) to the reaction resulted in complete inhibition of amido-4-methylcoumarylamide production, indicating successful isolation of proteasomes without the presence of significant amounts of other proteases.

Proteasome content analysis. For determining the proteasome content in the diaphragm muscle, a Western blot was performed by using an antibody against the 20S proteasome subunit C8 (MCP72; mouse monoclonal; 1:10,000; Affiniti, Gorinchem, The Netherlands). In short, frozen diaphragm samples were homogenized in ~200 µl ice-cold buffer (pH 7.5) containing (in mM) 50 Tris, 1 EDTA, 1 dithiothreitol, 1 PMSF, and protease inhibitor cocktail (Sigma) by means of a Dounce homogenizer. Homogenates were centrifuged at 10,000 g at 4°C for 10 min, and the protein concentration of the resulting supernatants was determined by using the Bio-Rad protein assay. Soluble proteins (10 µg) were subjected to routine Western blotting by using 10% polyacrylamide SDS gels and the MCP72 antibody. After being washed, the blot was incubated with a horseradish peroxidase-conjugated goat anti-mouse Ig (1:10,000; Pierce, Etten-Leur, The Netherlands). Chemiluminescence was performed by using an ECL Western-blotting analysis system (Amersham Biosciences, Roosendaal, The Netherlands). Protein bands were quantified by using optical densitometry software (GeneTools). An isolated proteasome fraction from a sham-operated rat was used as a positive control, and for the negative control a homogenate of a sham-operated rat was stained without addition of the primary (anti-C8) antibody. Equal loading was controlled by Coomassie blue staining.

Data treatment and statistical methods. Contractile properties of 82 diaphragm muscle single fibers from 14 CHF rats and 70 single fibers from 6 sham-operated rats were measured. These fibers expressed a single myosin heavy chain isoform, and contractile data were grouped per fiber type: slow, 2a, and 2x. Every rat was equally represented in the fiber-type groups. None of the fibers solely expressed myosin heavy chain 2b isoform. Fibers that coexpressed myosin heavy chain isoforms (n = 3 in CHF fibers and n = 4 in sham-operated rat fibers) were excluded from further analysis. Due to technical circumstances, in some fibers not all contractile parameters could be measured. Therefore, the number of single fibers analyzed differs per parameter.

Differences in animal characteristics, caspase-3, and proteasome activity between groups were analyzed with a Student's t-test. Differences in parameters describing single-fiber characteristics and contractile properties between groups were tested with one-way ANOVA, followed by the Student-Newman-Keuls post hoc test. Because of a limited amount of diaphragm tissue per rat, proteasome activity, caspase-3 activity, and Western-blotting studies were not based on the same rats, although there is extensive overlap. A probability level of P < 0.05 was considered significant unless otherwise indicated. Mean ± SE values are presented in text, tables, and figures.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
CHF indexes. All rats with infarct sizes >35% of the left ventricle (n = 14) showed clinical signs of severe heart failure, including pulmonary congestion and cardiomegaly, as supported by increased wet-to-dry lung-weight ratio and heart weight, respectively (Table 1). Elevated left-ventricular end-diastolic pressure, decreased left-ventricular peak systolic pressure, and decreased aortic pressures indicate impaired left-ventricular function (Table 1).


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  		Table 1. Animal characteristics

 
Maximum force per cross-sectional area. Single-fiber cross-sectional area did not differ significantly between fibers from CHF and sham-operated rats (1.0 ± 0.1 x 10–3 vs. 1.1 ± 0.1 x 10–3 mm2 for slow fibers; 2.5 ± 0.5 x 10–3 vs. 2.4 ± 0.5 x 10–3 mm2 for type 2a fibers, and 3.3 ± 0.3 x 10–3 vs. 3.5 ± 0.4 x 10–3 mm2 for type 2x fibers, respectively). Cross-sectional areas of slow fibers were significantly smaller compared with 2a and 2x fibers for both groups (P < 0.05). Maximum force per cross-sectional area of single fibers from the diaphragm of CHF rats was significantly reduced compared with fibers from sham-operated rats for all fiber types (P < 0.01, Fig. 1). Maximum force per cross-sectional area did not significantly differ between fiber types within groups (Fig. 1).


Figure 1
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  		Fig. 1. Maximum force per cross-sectional area (CSA) in slow, type 2a, and type 2x diaphragm muscle single fibers. Maximum force per CSA of single fibers from the diaphragm of congestive heart failure (CHF) rats was significantly reduced compared with sham-operated rats for all fiber types. Numbers above bars represent number of single fibers analyzed. Data are presented as means ± SE. *P < 0.01 vs. sham-operated group. Fmax, maximum force.

 
Diaphragm muscle fiber myosin heavy chain concentration. Myosin heavy chain concentration in single fibers from CHF rats was significantly lower than in sham-operated rats for all fiber types (P < 0.05, Fig. 2A).


Figure 2
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  		Fig. 2. A: myosin heavy chain concentration in slow, type 2a, and type 2x fibers from the diaphragm of CHF rats are decreased compared with sham-operated rats for all fiber types. B: myosin heavy chain content per half-sarcomere in slow, type 2a, and type 2x fibers from the diaphragm of CHF are reduced compared with sham-operated rats for all fiber types. Type 2x fibers contained more myosin heavy chain per half-sarcomere than slow fibers for both the CHF and sham-operated groups. Numbers above bars represent number of single fibers analyzed. C: representative silver-stained gel loaded with rat diaphragm homogenate, single fibers from the diaphragm of CHF and sham-operated rats, and myosin standards. Data are presented as means ± SE. *P < 0.05 vs. sham-operated group. {dagger}P < 0.001 compared with slow fibers within group. MHC, myosin heavy chain.

 
Diaphragm muscle fiber myosin heavy chain content per half-sarcomere. Myosin heavy chain content per half-sarcomere reflects the number of cross-bridges in parallel and was determined from myosin heavy chain concentration and half-sarcomere volume. Myosin heavy chain content per half-sarcomere is reduced in single fibers from CHF rats compared with sham-operated rats for all fiber types (P < 0.01, Fig. 2B). Type 2x fibers contain more myosin heavy chain per half-sarcomere than slow fibers (P < 0.001 for both CHF and sham-operated rats; Fig. 2). Figure 2C shows a representative silver-stained gel.

Maximum force per half-sarcomere myosin heavy chain content. To evaluate the effect of the number of cross-bridges in parallel on the maximum force per cross-sectional area, maximal force generated by single fibers was normalized to myosin heavy chain content per half-sarcomere volume. Maximum force per myosin heavy chain content per half-sarcomere was not significantly different between CHF and sham-operated diaphragm for all fiber types (Fig. 3). Maximum force per myosin heavy chain content per half-sarcomere was lower in slow fibers than in 2a and 2x fibers for both groups, but this difference did not reach significance (Fig. 3).


Figure 3
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  		Fig. 3. Single-fiber maximum force per half-sarcomere myosin heavy chain content in slow, type 2a, and type 2x fibers from the diaphragm of CHF and sham-operated rats. Maximum force normalized for half-sarcomere myosin heavy chain content provides an estimation for the average force generated per cross-bridge. Maximum force per half-sarcomere myosin heavy chain content was not significantly different between CHF and sham-operated diaphragm for all fiber types. Numbers above bars represent total number of single fibers analyzed. Data are presented as means ± SE.

 
Fraction of cross-bridges in strongly attached state during maximal activation. Maximum force also depends on the fraction of all available cross-bridges that are in the strongly attached state during maximal activation. The fraction of cross-bridges in the strongly attached state during maximal activation was not significantly different between diaphragm muscle single fibers from CHF and sham-operated rats (Fig. 4).


Figure 4
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  		Fig. 4. Fraction of strongly attached cross-bridges ({alpha}fs) at maximal activation. Fraction was not significantly different between slow, type 2a, and type 2x fibers from the diaphragm of CHF and sham-operated rats. Numbers above bars represent total number of single fibers analyzed. Data are presented as means ± SE.

 
Cross-bridge cycling kinetics. The ktr was slower in diaphragm muscle fibers from CHF rats than sham-operated rat fibers (P < 0.05, Fig. 5). As expected, ktr of type 2a and 2x fibers was significantly faster than in slow fibers in both groups (P < 0.01, Fig. 5). Figure 5B shows representative force redevelopment signals for the different fiber types.


Figure 5
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  		Fig. 5. Rate constant of force redevelopment (ktr) was used as a measure for cross-bridge cycling kinetics. A: ktr in slow, type 2a, and type 2x fibers from the diaphragm of CHF was slower than in diaphragm fibers from sham-operated rats. ktr at maximal activation was significantly faster in type 2a and 2x fibers than in slow fibers for both groups. Numbers above bars represent total number of single fibers analyzed. B: representative plots of force vs. time during the ktr protocol for slow fibers (B1), 2a fibers (B2) and 2x fibers (B3). After rapid detachment of all cross-bridges, force redevelops from 0 according to a first-order exponential curve. ktr value represents time constant of this curve. Data are presented as means ± SE. *P < 0.05 vs. sham group. {dagger}P < 0.01 compared with slow fibers within group.

 
Ca2+ sensitivity of force generation. The force-pCa curves for slow and 2x fibers of CHF rats are shifted rightward compared with sham-operated rats (Fig. 6). Consequently, the negative logarithm of pCa50 is significantly reduced for slow and type 2x diaphragm fibers from CHF rats compared with sham-operated rats (5.36 ± 0.05 vs. 5.73 ± 0.02, P < 0.01 for slow and 4.91 ± 0.06 vs. 5.16 ± 0.04, P < 0.05 for type 2x, respectively). The pCa50 of type 2x fibers was significantly lower compared with slow fibers in both groups (P < 0.001). Hill coefficients were not significantly different between fibers from CHF and sham-operated rats (1.9 ± 0.2 vs. 1.8 ± 0.2 for slow and 2.3 ± 0.2 vs. 2.6 ± 0.3 for type 2x, respectively). The Hill coefficients for type 2x fibers were significantly higher than for slow fibers in the sham group (P < 0.01). Due to a limited presence of 2a fibers in the rat diaphragm, we were unable to measure the force-pCa relation of a sufficient amount of 2a fibers in both CHF and sham-operated groups.


Figure 6
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  		Fig. 6. Force-pCa relation of single fibers from the diaphragm of CHF and sham-operated rats. Isometric force generated in response to incubation of single fibers with incremental Ca2+ concentrations was determined. Force-pCa relations of CHF diaphragm slow and 2x fibers are shifted to the right compared with sham-operated rats. Data are presented as means ± SE. pCa, negative log of Ca2+ concentration.

 
Caspase-3 activity. Caspase-3 activity of diaphragm homogenates was ~1.3-fold higher for CHF compared with sham-operated rats (P < 0.05, Fig. 7).


Figure 7
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  		Fig. 7. Caspase-3 activity in the diaphragm of CHF (n = 10) and sham-operated (n = 10) rats. Caspase-3 activity against the fluorogenic substrate N-acetyl-Asp-Glu-Val-Asp-7-amido-4-methylcoumarin (AMC) was higher in the diaphragm homogenates from CHF than from sham-operated rats. Data are presented as means ± SE. *P < 0.05 vs. sham-operated rats.

 
Proteasome activity and content. The proteasome activity against both LLVY and LLE was ~1.6-fold higher in diaphragm from CHF rats compared with sham-operated rats (P < 0.05, Fig. 8A). To investigate if changes in proteasome activity are the result of changes in the number of proteasomes, we determined the protein level of the C8 subunit of the 20S proteasome. Figure 8B demonstrates that the protein levels of the 20S proteasome C8 subunit were not significantly different between the diaphragm from CHF and sham-operated rats.


Figure 8
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  		Fig. 8. Proteasome activity and content in the diaphragm of CHF (n = 6) and sham-operated (n = 6) rats. A: proteasome activity against fluorogenic substrates succinyl-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin (LLVY) and N-carbenzoxy-Leu-Leu-Glu-7-amido-4-methylcoumarin (LLE) was higher in CHF than in sham-operated rats. B: subunit C8 protein content was determined in diaphragm homogenates by Western blotting and subsequent densitometric quantification of protein bands. Top: our analyses revealed no significantly different contents of subunit C8 in diaphragm of CHF and sham-operated rats. Bottom: representative C8 subunit immunoblot from diaphragm homogenates from CHF and sham-operated rats. Data are presented as means ± SE. *P < 0.05 vs. sham-operated rats.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The present study is the first to investigate the effects of CHF on diaphragm skinned single fiber function. The data from our study show that single fibers from the diaphragm of CHF rats generate less force per cross-sectional area for every fiber type. In these fibers, myosin heavy chain content per half-sarcomere was concomitantly reduced. Therefore, when normalized for myosin content per half-sarcomere, maximum force of diaphragm fibers was not different between CHF and sham-operated rats for every fiber type. In addition to reduced maximal force generation, CHF decreased Ca2+ sensitivity of force generation and slowed cross-bridge cycling kinetics in every fiber type. Finally, CHF enhanced caspase-3 and proteasome activity in the diaphragm.

Comparisons of control data with other studies. Single fiber-specific force in sham-operated rats was not different between fiber types, which is in line with previous publications from other groups (42, 51), although some controversy exists (3, 24). In agreement with the first report on diaphragm fiber-type differences in cross-bridge cycling kinetics (47), the present study shows that ktr of slow fibers is lower than for fast (2x and 2a) fibers. Differences in myosin heavy chain content per half-sarcomere, calcium sensitivity, and Hill coefficients between fiber types are also in accordance with previous studies on rat diaphragm fibers (8, 23, 25). However, absolute values of myosin content per half sarcomere and pCa50 in sham-operated diaphragm single fibers were slightly different from reports by other groups (23, 25, 51). These differences could possibly have arisen from the use of dissimilar compositions of activating solutions, animal race, and age. Nevertheless, it is unlikely that the discrepancies can explain the significant differences between sham-operated and CHF diaphragm fiber contractile properties presented in this study. Values of cross-sectional area and the fraction of all available cross-bridges that are in the strongly attached state ({alpha}fs) of sham-operated diaphragm fibers were comparable with results of previous studies on rat diaphragm fibers (22, 23).

Causes of impaired diaphragm muscle single-fiber contractility in CHF. Diaphragm muscle bundle isometric contractility has been evaluated in several animal models for CHF (29, 49, 50). However, no previous studies have investigated the contractile performance of diaphragm muscle single fibers in CHF. This study shows that maximal force-generating capacity of skinned single fibers from the diaphragm of CHF rats is reduced in all fiber types. This suggests that the reduced maximal force generation of the diaphragm in CHF patients (28) and animal models (29, 49, 50) is not merely attributable to a fiber-type shift but rather to a decrease of maximal force generation of individual fibers.

Maximal force (Fmax) generation of skinned muscle fibers is determined by the sum of forces generated by individual cross-bridges (6, 24). Therefore

Formula
where n is the number of available cross-bridges in parallel per half-sarcomere and fm is the mean force per attached cross-bridge in the force-generating state.

The reduced maximal force generation of the diaphragm muscle fibers from CHF rats should be reflected by a reduction in one or more of these determinants. Indeed, we showed that myosin heavy chain concentration was decreased in all fiber types in the diaphragm from CHF rats (Fig. 2A). Because fiber cross-sectional areas were not different between groups, this indicates that myosin heavy chain content per half-sarcomere was significantly reduced in CHF fibers compared with sham-operated rats (Fig. 2B). This reduction causes a proportional decrease in n. The {alpha}fs (Fig. 4) was not different between diaphragm fibers from CHF and sham-operated rats. Also, the fm, which can be derived from Fmax (Fig. 3) and {alpha}fs (see Eq. 1), did not differ between both groups. Therefore, the reduction in maximal force generation of diaphragm fibers from CHF rats can be mainly explained by the loss of myosin heavy chain content (n) in these fibers, although the involvement of other processes cannot be excluded.

A recent study (58) showed no differences of myosin heavy chain content between diaphragm homogenates of Dahl salt-sensitive rats with and without heart failure. Next to a different animal model of heart failure that was used in that study, another reason for dissimilar results lies in the difference in reliability of the methods that were used to measure myosin heavy chain content. Measuring myosin content in diaphragm homogenates is sensitive for differences in the amount of connective tissue and fiber-type distribution, whereas this variance is omitted in myosin content determinations on single fibers, as was done in the present study. Moreover, correcting myosin content for the total amount of protein in diaphragm homogenates underestimates a loss of myosin, because myosin constitutes a significant fraction of the total protein amount. Correcting myosin content for fiber volume in single fibers is therefore more accurate.

The most important findings of this study include decreased maximal force generation together with a loss of myosin in CHF diaphragm fibers. However, in vivo the diaphragm does not perform maximal isometric contractions during normal breathing but shortens against a certain load. Therefore, submaximal and kinetic parameters of single-fiber measurements provide more relevant physiological information. The rightward shift of the pCa-force curve displays that at a certain Ca2+ concentration, a smaller percentage of maximal force is generated in diaphragm fibers from CHF rats compared with sham-operated rats. In other words, Ca2+ sensitivity of force generation is reduced. This implies that force generation in the CHF diaphragm is even more affected at submaximal than at maximal activation, which is in line with a previously observed rightward shift of the force-frequency relation in CHF diaphragm bundles (52). The decreased rate constant of force redevelopment shows that cross-bridge cycling kinetics are slower in CHF diaphragm fibers compared with sham-operated fibers. This observation is in accordance with the decreased in vitro sliding velocities of myosin, isolated from the diaphragm of CHF rats, recently described by Coirault et al. (12). However, by using an in vitro motility assay, Coirault et al. could not exclude a possible effect of fiber-type shift on the decreased sliding velocities of myosin. Our single-fiber data unambiguously show that slower cross-bridge kinetics is an intrinsic phenomenon of all fiber types in the CHF diaphragm.

The causes of slower cross-bridge cycling kinetics and decreased Ca2+ sensitivity in the CHF diaphragm remain speculative. Previous studies showed that increased filament-lattice spacing results in slower rates of tension redevelopment (33) and decreased Ca2+ sensitivity (26). Indeed, in our study myosin content per half-sarcomere is reduced in the CHF diaphragm (Fig. 2) without alterations of cross-sectional area, indicating increased filament-lattice spacing. Thus myosin loss in CHF diaphragm fibers could not only result in decreased maximal force generation but may also affect submaximal and kinetic parameters of diaphragm contractility. Besides, modification of the myosin head or the Ca2+-binding protein complex troponin could also account for slower cross-bridge kinetics and reduced Ca2+ sensitivity (41, 43, 45). For example, in their in vitro motility study, Coirault et al. (12) showed that the decreased sliding velocities of myosin are associated with increased levels of myosin oxidation in the diaphragm of CHF rats. However, further studies on filament-lattice spacing and posttranslational modifications of contractile proteins are needed to establish the precise origin of slower cross-bridge kinetics and decreased Ca2+ sensitivity in CHF diaphragm fibers.

Pathways toward a loss of myosin heavy chain content in CHF. The loss of myosin heavy chain in the CHF diaphragm suggests an imbalance between the synthesis and degradation of this contractile protein. In peripheral skeletal muscles of CHF patients, rates of myosin heavy chain synthesis were not different from control patients, whereas the decrease in muscle strength was closely related to the loss of myosin (57). This suggests that enhanced degradation of myosin needs to be involved. Indeed, recent studies have indicated that the degradation of myosin plays an important role in diseases associated with diaphragm (39) and peripheral muscle wasting (1, 30), including CHF (46). It is well recognized that myofibrillar proteins such as myosin are degraded by the ubiquitin-proteasome pathway (34, 36, 55). The present study shows that the activity of the ubiquitin-proteasome pathway is increased in the CHF diaphragm. More specifically, these data indicate that the proteolytic activity per proteasome is increased rather than the number of proteasomes. Thus in the CHF diaphragm, the loss of myosin heavy chain is associated with an increased activity of the ubiquitin-proteasome pathway.

Because the ubiquitin-proteasome pathway is unable to degrade intact actomyosin complexes (48), preceding cleavage of these proteins from the sarcomere is necessary for subsequent degradation. The exact mechanisms for cleaving contractile proteins from the sarcomere are unknown, but an important role is ascribed to proteolytic enzymes, such as calpains (2, 60) and caspase-3 (18, 53). Previously, a twofold increase of calpain activity in the diaphragm of CHF rats has been reported (17). This study demonstrates that caspase-3 activity is enhanced in the CHF diaphragm and is associated with loss of myosin and activated proteasomes. However, further studies are needed to elucidate the exact role for caspase-3 activation in CHF diaphragm weakness.

Conclusion. This study shows diaphragm weakness at the single-fiber level in a myocardial infarction model of CHF. Maximal force generation of diaphragm muscle single fibers from CHF rats is reduced and can be explained by a decreased myosin heavy chain content in all fiber types. Additionally, cross-bridge kinetics are slower and Ca2+ sensitivity is decreased in the diaphragm fibers of CHF rats. Finally, CHF diaphragm weakness and the loss of myosin heavy chain are associated with an increased activation of the ubiquitin proteasome pathway and its upstream proteolytic enzyme caspase-3.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported by an unrestricted educational grant from Novartis.


   ACKNOWLEDGMENTS
 
We are indebted to L. Ennen and M. Linkels (Radboud University Nijmegen Medical Centre, Nijmegen, The Netherlands) for expert technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: H. W. H. van Hees, Dept. of Pulmonary Diseases - 454, Univ. Medical Centre Nijmegen, PO Box 9101, 6500 HB Nijmegen, The Netherlands (e-mail: H.vanhees{at}long.umcn.nl)

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
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Acharyya S, Ladner KJ, Nelsen LL, Damrauer J, Reiser PJ, Swoap S, Guttridge DC. Cancer cachexia is regulated by selective targeting of skeletal muscle gene products. J Clin Invest 114: 370–378, 2004.[CrossRef][ISI][Medline]
  2. Bartoli M, Richard I. Calpains in muscle wasting. Int J Biochem Cell Biol 37: 2115–2133, 2005.[CrossRef][ISI][Medline]
  3. Bottinelli R, Canepari M, Reggiani C, Stienen GJ. Myofibrillar ATPase activity during isometric contraction and isomyosin composition in rat single skinned muscle fibres. J Physiol 481: 663–675, 1994.[Abstract/Free Full Text]
  4. Brenner B. Muscle mechanics II: skinned muscle fibers. In: Current Methods In Muscle Physiology: Advantages, Problems and Limitations, edited by Sugi H. New York: Oxford University Press, 1998, p. 33–69.
  5. Brenner B. Technique for stabilizing the striation pattern in maximally calcium-activated skinned rabbit psoas fibers. Biophys J 41: 99–102, 1983.[ISI][Medline]
  6. Brenner B. Effect of Ca2+ on cross-bridge turnover kinetics in skinned single rabbit psoas fibers: implications for regulation of muscle contraction. Proc Natl Acad Sci USA 85: 3265–3269, 1988.[Abstract/Free Full Text]
  7. Brenner B, Eisenberg E. Rate of force generation in muscle: correlation with actomyosin ATPase activity in solution. Proc Natl Acad Sci USA 83: 3542–3546, 1986.[Abstract/Free Full Text]
  8. Brotto MA, Biesiadecki BJ, Brotto LS, Nosek TM, Jin JP. Coupled expression of troponin T and troponin I isoforms in single skeletal muscle fibers correlates with contractility. Am J Physiol Cell Physiol 290: C567–C576, 2006.[Abstract/Free Full Text]
  9. Burkholder TJ, Lieber RL. Sarcomere length operating range of vertebrate muscles during movement. J Exp Biol 204: 1529–1536, 2001.[Abstract]
  10. Chamberlain JS. Cachexia in cancer–zeroing in on myosin. N Engl J Med 351: 2124–2125, 2004.[Free Full Text]
  11. Cicoira M, Zanolla L, Franceschini L, Rossi A, Golia G, Zamboni M, Tosoni P, Zardini P. Skeletal muscle mass independently predicts peak oxygen consumption and ventilatory response during exercise in noncachectic patients with chronic heart failure. J Am Coll Cardiol 37: 2080–2085, 2001.[Abstract/Free Full Text]
  12. Coirault C, Guellich A, Barbry T, Samuel JL, Riou B, Lecarpentier Y. Oxidative stress of myosin contributes to skeletal muscle dysfunction in rats with chronic heart failure. Am J Physiol Heart Circ Physiol 292: H1009–H1017, 2007.[Abstract/Free Full Text]
  13. Craiu A, Gaczynska M, Akopian T, Gramm CF, Fenteany G, Goldberg AL, Rock KL. Lactacystin and clasto-lactacystin beta-lactone modify multiple proteasome beta-subunits and inhibit intracellular protein degradation and major histocompatibility complex class I antigen presentation. J Biol Chem 272: 13437–13445, 1997.[Abstract/Free Full Text]
  14. De Sousa E, Veksler V, Bigard X, Mateo P, Serrurier B, Ventura-Clapier R. Dual influence of disease and increased load on diaphragm muscle in heart failure. J Mol Cell Cardiol 33: 699–710, 2001.[CrossRef][ISI][Medline]
  15. Dominguez JF, Howell S. Compartmental analysis of steady-state diaphragm Ca2+ kinetics in chronic congestive heart failure. Cell Calcium 33: 163–174, 2003.[CrossRef][ISI][Medline]
  16. Du J, Wang X, Miereles C, Bailey JL, Debigare R, Zheng B, Price SR, Mitch WE. Activation of caspase-3 is an initial step triggering accelerated muscle proteolysis in catabolic conditions. J Clin Invest 113: 115–123, 2004.[CrossRef][ISI][Medline]
  17. Filippatos GS, Anker SD, Kremastinos DT. Pathophysiology of peripheral muscle wasting in cardiac cachexia. Curr Opin Clin Nutr Metab Care 8: 249–254, 2005.[ISI][Medline]
  18. Ford LE, Huxley AF, Simmons RM. The relation between stiffness and filament overlap in stimulated frog muscle fibres. J Physiol 311: 219–249, 1981.[Abstract/Free Full Text]
  19. Franciosa JA, Park M, Levine TB. Lack of correlation between exercise capacity and indexes of resting left ventricular performance in heart failure. Am J Cardiol 47: 33–39, 1981.[CrossRef][ISI][Medline]
  20. Geiger PC, Cody MJ, Han YS, Hunter LW, Zhan WZ, Sieck GC. Effects of hypothyroidism on maximum specific force in rat diaphragm muscle fibers. J Appl Physiol 92: 1506–1514, 2002.[Abstract/Free Full Text]
  21. Geiger PC, Cody MJ, Macken RL, Bayrd ME, Sieck GC. Effect of unilateral denervation on maximum specific force in rat diaphragm muscle fibers. J Appl Physiol 90: 1196–1204, 2001.[Abstract/Free Full Text]
  22. Geiger PC, Cody MJ, Macken RL, Sieck GC. Maximum specific force depends on myosin heavy chain content in rat diaphragm muscle fibers. J Appl Physiol 89: 695–703, 2000.[Abstract/Free Full Text]
  23. Geiger PC, Cody MJ, Sieck GC. Force-calcium relationship depends on myosin heavy chain and troponin isoforms in rat diaphragm muscle fibers. J Appl Physiol 87: 1894–1900, 1999.[Abstract/Free Full Text]
  24. Godt RE, Maughan DW. Influence of osmotic compression on calcium activation and tension in skinned muscle fibers of the rabbit. Pflügers Arch 391: 334–337, 1981.[ISI][Medline]
  25. Howell S, Maarek JM, Fournier M, Sullivan K, Zhan WZ, Sieck GC. Congestive heart failure: differential adaptation of the diaphragm and latissimus dorsi. J Appl Physiol 79: 389–397, 1995.[Abstract/Free Full Text]
  26. Hughes PD, Polkey MI, Harrus ML, Coats AJ, Moxham J, Green M. Diaphragm strength in chronic heart failure. Am J Respir Crit Care Med 160: 529–534, 1999.[Abstract/Free Full Text]
  27. Lecarpentier Y, Coirault C, Langeron O, Blanc FX, Salmeron S, Attal P, Riou B, Chemla D. Impaired load dependence of diaphragm relaxation during congestive heart failure in the rabbit. J Appl Physiol 87: 1339–1345, 1999.[Abstract/Free Full Text]
  28. Lecker SH, Jagoe RT, Gilbert A, Gomes M, Baracos V, Bailey J, Price SR, Mitch WE, Goldberg AL. Multiple types of skeletal muscle atrophy involve a common program of changes in gene expression. FASEB J 18: 39–51, 2004.[Abstract/Free Full Text]
  29. Loot AE, Roks AJ, Henning RH, Tio RA, Suurmeijer AJ, Boomsma F, van Gilst WH. Angiotensin-(1–7) attenuates the development of heart failure after myocardial infarction in rats. Circulation 105: 1548–1550, 2002.[Abstract/Free Full Text]
  30. Mancini DM, Henson D, LaManca J, Levine S. Respiratory muscle function and dyspnea in patients with chronic congestive heart failure. Circulation 86: 909–918, 1992.[Abstract/Free Full Text]
  31. McDonald KS, Wolff MR, Moss RL. Sarcomere length dependence of the rate of tension redevelopment and submaximal tension in rat and rabbit skinned skeletal muscle fibres. J Physiol 501: 607–621, 1997.[Abstract/Free Full Text]
  32. McKinnell IW, Rudnicki MA. Molecular mechanisms of muscle atrophy. Cell 119: 907–910, 2004.[CrossRef][ISI][Medline]
  33. Meyer FJ, Borst MM, Zugck C, Kirschke A, Schellberg D, Kubler W, Haass M. Respiratory muscle dysfunction in congestive heart failure: clinical correlation and prognostic significance. Circulation 103: 2153–2158, 2001.[Abstract/Free Full Text]
  34. Mitch WE, Goldberg AL. Mechanisms of muscle wasting. The role of the ubiquitin-proteasome pathway. N Engl J Med 335: 1897–1905, 1996.[Free Full Text]
  35. Oakley BR, Kirsch DR, Morris NR. A simplified ultrasensitive silver stain for detecting proteins in polyacrylamide gels. Anal Biochem 105: 361–363, 1980.[CrossRef][ISI][Medline]
  36. Okita K, Yonezawa K, Nishijima H, Hanada A, Ohtsubo M, Kohya T, Murakami T, Kitabatake A. Skeletal muscle metabolism limits exercise capacity in patients with chronic heart failure. Circulation 98: 1886–1891, 1998.[Abstract/Free Full Text]
  37. Ottenheijm CA, Heunks LM, Li YP, Jin B, Minnaard R, van Hees HW, Dekhuijzen PN. Activation of the ubiquitin-proteasome pathway in the diaphragm in chronic obstructive pulmonary disease. Am J Respir Crit Care Med 174: 997–1002, 2006.[Abstract/Free Full Text]
  38. Ottenheijm CA, Heunks LM, Sieck GC, Zhan WZ, Jansen SM, Degens H, de BT, Dekhuijzen PN. Diaphragm dysfunction in chronic obstructive pulmonary disease. Am J Respir Crit Care Med 172: 200–205, 2005.[Abstract/Free Full Text]
  39. Park HS, Gong BJ, Tao T. A disulfide crosslink between Cys98 of troponin-C and Cys133 of troponin-I abolishes the activity of rabbit skeletal troponin. Biophys J 66: 2062–2065, 1994.[ISI][Medline]
  40. Patterson MF, Stephenson GM, Stephenson DG. Denervation produces different single fiber phenotypes in fast- and slow-twitch hindlimb muscles of the rat. Am J Physiol Cell Physiol 291: C518–C528, 2006.[CrossRef][ISI][Medline]
  41. Perkins WJ, Han YS, Sieck GC. Skeletal muscle force and actomyosin ATPase activity reduced by nitric oxide donor. J Appl Physiol 83: 1326–1332, 1997.[Abstract/Free Full Text]
  42. Pfeffer MA, Pfeffer JM, Fishbein MC, Fletcher PJ, Spadaro J, Kloner RA, Braunwald E. Myocardial infarct size and ventricular function in rats. Circ Res 44: 503–512, 1979.[Abstract/Free Full Text]
  43. Root DD, Cheung P, Reisler E. Catalytic cooperativity induced by SH1 labeling of myosin filaments. Biochemistry 30: 286–294, 1991.[CrossRef][Medline]
  44. Schulze PC, Fang J, Kassik KA, Gannon J, Cupesi M, MacGillivray C, Lee RT, Rosenthal N. Transgenic overexpression of locally acting insulin-like growth factor-1 inhibits ubiquitin-mediated muscle atrophy in chronic left-ventricular dysfunction. Circ Res 97: 418–426, 2005.[Abstract/Free Full Text]
  45. Sieck GC, Prakash YS. Cross-bridge kinetics in respiratory muscles. Eur Respir J 10: 2147–2158, 1997.[Abstract]
  46. Solomon V, Baracos V, Sarraf P, Goldberg AL. Rates of ubiquitin conjugation increase when muscles atrophy, largely through activation of the N-end rule pathway. Proc Natl Acad Sci USA 95: 12602–12607, 1998.[Abstract/Free Full Text]
  47. Stassijns G, Gayan-Ramirez G, De Leyn P, de B, V, Dom R, Lysens R, Decramer M. Effects of dilated cardiomyopathy on the diaphragm in the Syrian hamster. Eur Respir J 13: 391–397, 1999.[Abstract]
  48. Supinski G, DiMarco A, Dibner-Dunlap M. Alterations in diaphragm strength and fatiguability in congestive heart failure. J Appl Physiol 76: 2707–2713, 1994.[Abstract/Free Full Text]
  49. Supinski G, Nethery D, Nosek TM, Callahan LA, Stofan D, DiMarco A. Endotoxin administration alters the force vs. pCa relationship of skeletal muscle fibers. Am J Physiol Regul Integr Comp Physiol 278: R891–R896, 2000.[Abstract/Free Full Text]
  50. Supinski GS, Callahan LA. Diaphragmatic free radical generation increases in an animal model of heart failure. J Appl Physiol 99: 1078–1084, 2005.[Abstract/Free Full Text]
  51. Supinski GS, Callahan LA. Caspase activation contributes to endotoxin-induced diaphragm weakness. J Appl Physiol 100: 1770–1777, 2006.[Abstract/Free Full Text]
  52. Sweeney HL, Corteselli SA, Kushmerick MJ. Measurements on permeabilized skeletal muscle fibers during continuous activation. Am J Physiol Cell Physiol 252: C575–C580, 1987.[Abstract/Free Full Text]
  53. Taillandier D, Combaret L, Pouch MN, Samuels SE, Bechet D, Attaix D. The role of ubiquitin-proteasome-dependent proteolysis in the remodelling of skeletal muscle. Proc Nutr Soc 63: 357–361, 2004.[CrossRef][ISI][Medline]
  54. Thomas GD, Zhang W, Victor RG. Impaired modulation of sympathetic vasoconstriction in contracting skeletal muscle of rats with chronic myocardial infarctions: role of oxidative stress. Circ Res 88: 816–823, 2001.[Abstract/Free Full Text]
  55. Toth MJ, Matthews DE, Ades PA, Tischler MD, Van Buren P, Previs M, Le Winter MM. Skeletal muscle myofibrillar protein metabolism in heart failure: Relationship to immune activation and functional capacity. Am J Physiol Endocrinol Metab 288: E883–E891, 2005.[Abstract/Free Full Text]
  56. Toth MJ, Palmer BM, LeWinter MM. Effect of heart failure on skeletal muscle myofibrillar protein content, isoform expression and calcium sensitivity. Int J Cardiol 107: 211–219, 2006.[CrossRef][ISI][Medline]
  57. Vescovo G, Serafini F, Dalla LL, Leprotti C, Facchin L, Tenderini P, Ambrosio GB. Skeletal muscle myosin heavy chains in heart failure: correlation between magnitude of the isozyme shift, exercise capacity, and gas exchange measurements. Am Heart J 135: 130–137, 1998.[CrossRef][ISI][Medline]
  58. Williams AB, Decourten-Myers GM, Fischer JE, Luo G, Sun X, Hasselgren PO. Sepsis stimulates release of myofilaments in skeletal muscle by a calcium-dependent mechanism. FASEB J 13: 1435–1443, 1999.[Abstract/Free Full Text]
  59. Zuurbier CJ, Heslinga JW, Lee-de Groot MB, van der Laarse WJ. Mean sarcomere length-force relationship of rat muscle fibre bundles. J Biomech 28: 83–87, 1995.[CrossRef][ISI][Medline]



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