Transgenic overexpression of γ-cytoplasmic actin protects against eccentric contraction-induced force loss in mdx mice
© Baltgalvis et al; licensee BioMed Central Ltd. 2011
Received: 25 June 2011
Accepted: 13 October 2011
Published: 13 October 2011
γ-cytoplasmic (γ-cyto) actin levels are elevated in dystrophin-deficient mdx mouse skeletal muscle. The purpose of this study was to determine whether further elevation of γ-cyto actin levels improve or exacerbate the dystrophic phenotype of mdx mice.
We transgenically overexpressed γ-cyto actin, specifically in skeletal muscle of mdx mice (mdx-TG), and compared skeletal muscle pathology and force-generating capacity between mdx and mdx-TG mice at different ages. We investigated the mechanism by which γ-cyto actin provides protection from force loss by studying the role of calcium channels and stretch-activated channels in isolated skeletal muscles and muscle fibers. Analysis of variance or independent t-tests were used to detect statistical differences between groups.
Levels of γ-cyto actin in mdx-TG skeletal muscle were elevated 200-fold compared to mdx skeletal muscle and incorporated into thin filaments. Overexpression of γ-cyto actin had little effect on most parameters of mdx muscle pathology. However, γ-cyto actin provided statistically significant protection against force loss during eccentric contractions. Store-operated calcium entry across the sarcolemma did not differ between mdx fibers compared to wild-type fibers. Additionally, the omission of extracellular calcium or the addition of streptomycin to block stretch-activated channels did not improve the force-generating capacity of isolated extensor digitorum longus muscles from mdx mice during eccentric contractions.
The data presented in this study indicate that upregulation of γ-cyto actin in dystrophic skeletal muscle can attenuate force loss during eccentric contractions and that the mechanism is independent of activation of stretch-activated channels and the accumulation of extracellular calcium.
Keywordsstretch-activated channels calcium skeletal muscle injury dystrophin costamere
Duchenne muscular dystrophy
Dulbecco's modified Eagle's medium
extensor digitorum longus
flexor digitorum brevis
store-operated calcium entry
stromal interacting molecule 1
transient receptor potential
Duchenne muscular dystrophy (DMD) is a severe muscle-wasting disease caused by mutations in the dystrophin gene. Dystrophin localizes primarily to costameres, where it links the cortical actin cytoskeleton to the sarcolemma and the extracellular matrix . It is part of the dystrophin-glycoprotein complex (DGC), a large oligomeric complex of proteins thought primarily to stabilize the sarcolemma during muscle contraction. In the absence of a functional dystrophin protein, myofibers are susceptible to injury, leading to muscle degeneration, inflammation and fibrosis.
Dystrophin binds to filamentous actin with high affinity via two distinct actin-binding domains: an N-terminal tandem calponin homology domain and a region of basic spectrin-like repeats in the middle rod domain . In skeletal muscle, dystrophin forms a mechanically strong link with the cortical actin cytoskeleton, which is composed of γ-cytoplasmic (γ-cyto) and β-cytoplasmic actins [2, 3]. The tight association between dystrophin and actin has been demonstrated with inside-out peeled sarcolemma from wild-type (WT) myofibers in which γ-cyto actin was retained in a riblike costameric pattern but absent from the dystrophin-deficient mdx mouse sarcolemma . The importance of dystrophin-actin binding in vivo is also evident in transgenic mdx mice that express various dystrophin deletion constructs. Despite restoration of DGC members at the sarcolemma, at least one actin-binding domain is essential for partial or complete rescue of the dystrophic phenotype in transgenic mdx mice [4–6].
A robust phenotype of the mdx mouse is increased susceptibility to injury following eccentric muscle contractions [7, 8], a standard procedure to determine the efficacy of a drug or therapy in preclinical trials . Force loss following eccentric contractions depends on the number of contractions, speed of contraction, overall length change, muscle type and age of the mice. Overall force loss in WT mice ranges from 2% to 29%, but force loss can range from 27% to 69% in mdx mice . The mechanism of force loss following eccentric contractions in healthy muscle has been investigated. The majority of force loss following eccentric contractions in an in vivo rodent model was due to excitation-contraction uncoupling, and about 25% of the force loss can be attributed to physical damage . After the initial bout of injury, the loss of contractile protein accounts for strength loss during 14 to 28 days of recovery. It is unknown whether excitation-contraction uncoupling, physical damage, loss of contractile protein or a combination of different mechanisms explains the rapid force loss in mdx mice following eccentric contractions.
Recent studies have suggested that stretch-activated cation channels (calcium or sodium) at the sarcolemma contribute to force loss during eccentric contractions [12–15]. The removal of calcium in the muscle-bathing media or the addition of streptomycin to block stretch-activated channels has been shown to reverse or attenuate force loss during injurious muscle contractions in dystrophic muscle [12, 15–17]. Store-operated calcium entry (SOCE) is an important mechanism whereby calcium is brought into muscle cells during sarcoplasmic reticulum (SR) calcium depletion . It has been proposed that SOCE occurs through either (1) an interaction between transient receptor potential C (TRPC) channels and either the ryanodine receptor or inositol trisphosphate receptors or (2) the STIM1/Orai1 pathway . TRP channels are plasma membrane channels that vary in cation selectivity and sensitivity to mechanical stress. While TRP gene transcripts are generally in low abundance in skeletal muscle compared to some other tissues, TRPC3 is the most abundant, with TRP vanilloid 3 (TRPV3), TRPV4, TRPV6, TRP melastatin 3 (TRPM3), TRPM4 and TRPM7 also present . In fact, the overexpression of dominant-negative TRPC3 or TRPV2 channels in muscle ameliorates pathological changes [13, 21, 22], and TRPC3 overexpression in WT mice is sufficient to induce muscular dystrophy . mdx mice that overexpress a dominant-negative TRPV2 channel are also protected from extreme force loss following eccentric muscle contractions .
Although γ-cyto actin comprises only 1/4, 000th of the actin population in skeletal muscle , it is essential for the maintenance of normal muscle function in adults . We previously reported that γ-cyto actin levels are elevated fivefold in dystrophin-deficient skeletal muscle [23, 25, 26] and that muscle-specific ablation of γ-cyto actin does not exacerbate the dystrophic condition . As many costameric components are actin-binding proteins, it has been proposed that this increased γ-cyto actin expression may contribute to a compensatory remodeling response to reinforce the compromised sarcolemma in mdx mice . Understanding the function of increased γ-cyto actin in dystrophic muscle may identify a unique pharmacological target for the treatment of muscular dystrophies. The purpose of this study was to determine whether further elevation of γ-cyto actin levels improves or exacerbates the dystrophic phenotype of mdx mice. We generated a line of transgenic mdx mice (mdx-TG) that expressed γ-cyto actin 200-fold above mdx γ-cyto actin levels and showed substantial incorporation of γ-cyto actin into thin filaments. Interestingly, mdx-TG mice showed significantly attenuated force loss during eccentric muscle contractions through a mechanism independent of extracellular calcium and stretch-activated channels.
Generation of transgenic mice
Generation of Actg1-TG mice has been previously described . To generate dystrophin-deficient mdx-TG mice carrying the Actg1 transgene, Actg1-TG male mice were crossed with mdx females. TG-positive males were subsequently crossed with mdx females to generate mdx-TG mice. Animals were housed and treated in accordance with the standards set by the Institutional Animal Care and Use Committees at the University of Minnesota and the University of Wisconsin.
Determination of γ-cytoplasmic actin concentration in skeletal muscle
Known amounts of purified bovine brain actin  and SDS extracts from mdx-TG gastrocnemius or quadriceps femoris muscles were run side-by-side on a 3% to 12% SDS-polyacrylamide gel and transferred to nitrocellulose membrane. Two γ-cyto actin antibodies (monoclonal mouse 2 to 4 and polyclonal affinity-purified rabbit 7577) were used for Western blot analysis, each at a 1:1, 000 dilution. To determine the concentration of actin (in micrograms) in the SDS extracts, a concentration standard curve was derived from the known values of purified bovine γ-cyto actin using the densitometry and concentration standards feature in the LI-COR Odyssey Application Software version 2.1.12 (LI-COR Biosciences, Lincoln, NE, USA).
Skeletal muscle histology and confocal microscopy
Tibialis anterior (TA) and gastrocnemius muscles from WT, mdx and mdx-TG mice were frozen in melting isopentane and mounted in O.C.T. mounting medium for cryosectioning . Ten-micrometer sections were stained with hematoxylin and eosin. Centrally nucleated fibers (CNFs) were counted on whole sections and expressed as a percentage of the total number of fibers.
Isolation of myofibrils and confocal microscopy were both performed as previously described . Cross-sections of gastrocnemius muscles or myofibrils were stained using the following antibodies: affinity-purified γ-cyto actin rabbit 7577 (1:75 dilution; Sigma, St Louis, MO, USA), fast myosin heavy chain clone MY-32 (1:100 dilution; Sigma) and laminin α-2 rat clone 4H8-2 (1:200 dilution; Sigma). All images were obtained using a FluoView FV1000 single-photon confocal microscope equipped with a PlanAPO 60×/1.40 na oil immersion lens objective, a UPlan Fluorite N 1.30 NA 40× oil immersion lens objective and a UPLSAPO 0.40 NA 10× lens objective (all from Olympus America Inc, Melville, NY, USA). Images were collected with Olympus FluoView FV1000 version 1.7b software and assembled and processed using ImageJ software (National Institutes of Health, Bethesda, MD, USA) and Adobe Photoshop version 8.0 software (Adobe Systems Inc, San Jose, CA, USA).
Serum creatine kinase analysis
Serum was collected and stored at -80°C until analysis. Ten microliters of serum were applied to a VITROS DT60 II MicroSlide (Ortho-Clinical Diagnostics, Inc, Rochester, NY, USA) and creatine kinase (CK) activity measured using a Kodak Ektachem DT60 Analyzer (Eastman Kodak Co, Rochester, NY, USA) and a VITROS DTSC II Module (Ortho-Clinical Diagnostics, Inc).
Ex vivo contractility measurements and eccentric injury protocol
The extensor digitorum longus (EDL) muscles were assessed for contractile function and susceptibility to eccentric contraction-induced injury as previously described [26, 28]. Age-matched male mdx and mdx-TG mice (n = 5 or 6 per group) were anesthetized with sodium pentobarbital (100 mg/kg body weight). Their muscles were dissected and mounted onto a dual-mode muscle lever system (model 300B-LR Dual-Mode Lever Arm; Aurora Scientific Inc, Aurora, ON, Canada) with 4-0 suture in a 1.5-ml bath assembly filled with Krebs-Ringer bicarbonate buffer that was maintained at 25°C and perfused with 95% O2. The muscles were adjusted to their anatomic optimal length (Lo) based on resting tension of 0.4 g, and then muscle length was measured. The muscles remained quiescent in the bath for ten minutes. Passive stiffness, twitch force (Pt), tetanic force (Po) and active stiffness were measured prior to initiating the eccentric injury protocol. First, passive stiffness of the muscle was determined by stretching the muscle sinusoidally from 97.5% Lo to 102.5% Lo at 0.5 Hz while measuring the resulting force [29, 30]. Two twitches (Pt) separated by 30-second intervals were elicited, and the resulting force was measured. Maximal isometric tetanic contractions (Po) were obtained by stimulating muscles for 400 ms at 180 Hz and 150 V (Grass S48 Square Pulse Stimulator; Grass Technologies, West Warwick, RI, USA) delivered through a SIU-V stimulus isolation unit (Grass Technologies). Two minutes later a second tetanic isometric contraction was elicited, and at peak force a sinusoidal length oscillation of 0.01% Lo at 500 Hz was imposed to determine active stiffness, an indirect marker of strong-binding myosin [29, 30]. Two minutes after that an injury protocol consisting of five to ten eccentric contractions was begun. For these contractions, muscles were passively shortened from Lo to 0.95 Lo over 3 seconds, stimulated tetanically for 200 ms as the muscle lengthened to 1.05 Lo at 0.5 Lo/second and then passively returned to Lo. Each eccentric contraction was separated by three minutes of rest. The degree of injury was calculated as the percentage change in eccentric force production from the first to the last eccentric contraction. Specific force was calculated by dividing isometric force by the anatomical cross-sectional area of the muscle. The cross-sectional area of the muscle was calculated by dividing the mass of the muscle by the product of fiber length (Lf) (Lo × 0.44) and the density of muscle (1.06 g/mm3). Contractility and injury protocols were followed to study both the right and left sides and averaged as a single data point for each mouse when comparing mdx and mdx-TG mice.
To determine the effects of extracellular calcium and stretch-activated channels on the susceptibility to eccentric contraction-induced injury, three different conditions were tested, all using male mdx mice, with the contralateral muscle from the same mouse used as a control (n = 3 to 5 per group). In the first condition, Krebs-Ringer bicarbonate buffer was made without CaCl2 but replaced with the same molar ratio of MgCl2 (that is, calcium-free). In the second condition, 1 mM ethylene glycol tetraacetic acid (EGTA) was added to the calcium-free buffer. Pilot work revealed that EGTA blunted isometric force production, so buffer containing 1 mM EGTA was introduced only after the preinjury contractile measurements were completed. The muscle remained quiescent for ten minutes after the addition of the calcium-free buffer with EGTA before the eccentric contraction protocol was begun. In the third condition, 2 mM streptomycin was added to Krebs-Ringer bicarbonate buffer for the entire protocol. Pilot work revealed a greater effect on postinjury isometric force production with the 2 mM concentration instead of the previously reported 0.2 mM .
Flexor digitorum brevis myofiber isolation
Flexor digitorum brevis (FDB) fibers were isolated from WT and mdx mice as previously described [31–33]. The right and left FDB muscles were dissected from WT, mdx and mdx-TG mice and placed in 5 ml of sterile physiological rodent saline (138 mM NaCl, 2.7 mM KCl, 1.8 mM CaCl2, 1.06 mM MgCl2, 12.4 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 5.6 mM glucose, pH 7.3). The muscles were then transferred to a Petri dish containing 2 ml of media (DMEM, 10% fetal bovine serum, and 0.5% penicillin/streptomycin) containing 2 mg/ml collagenase. Following this step, they were incubated at 37°C (5% CO2) for three hours and then gently triturated in fresh media and allowed to settle at 1 × g. All but 1 ml of the media was aspirated, and the fiber pellet was gently resuspended. Next the fibers were placed into six 35-mm Petri dishes, each containing a glass coverslip coated with 1 μg of laminin supplemented with 2 ml of media. The fibers were allowed to adhere to the coverslips overnight at 37°C. They were analyzed for SOCE within 24 to 48 hours after dissection.
Store-operated calcium entry
SOCE was measured in FDB fibers as previously described [22, 34]. Coverslips containing adhered FDB fibers were briefly washed in physiological saline solution (PSS) (130 mM NaCl, 5.6 mM KCl, 1 mM MgCl2, 1.7 mM CaCl2, 11 mM glucose, 10 mM HEPES, pH 7.4). Indo-1 AM (Invitrogen, Carlsbad, CA, USA) was mixed 1:1 with Pluronic F-127 solution (Invitrogen) and added to myofibers at a final concentration of 1 μM. Myofibers were loaded with dye for 30 minutes at 37°C and washed with PSS for 15 minutes at 37°C. Instrumentation and recording of the calcium signal was performed as previously described . Myofibers were perfused with PSS at a rate of 1 to 2 ml/minute for two minutes before being moved to calcium-free PSS. The calcium-free PSS did not contain CaCl2, had a MgCl2 concentration of 2 mM, contained 1 mM EGTA and contained 30 μM cyclopiazonic acid (CPA) to block SR calcium-ATPase. Calcium-free PSS was perfused for four to eight minutes before 5 mM caffeine was added to release calcium from the SR. After the indo-1 ratio had returned to baseline (8 to 12 minutes), the muscle was perfused with PSS and SOCE was calculated as the difference in the minimum and maximum indo-1 ratios averaged over a 30-second period. During the recording of the indo-1 signal, 50 μM N-benzyl-p-toluene sulfonamide (BTS) was added to all solutions to prevent myofiber contractions.
Two-way analysis of variance (ANOVA) was used to assess differences in CNFs and serum CK with mouse strain (WT vs mdx vs mdx-TG) and age as the factors. One-way ANOVA was used to compare intracellular calcium concentration ([Ca2+]i) between WT, mdx and mdx-TG mice. Post hoc analyses were performed using the Holm-Sidak test. t-tests were used to analyze differences between all other variables. Data are reported as means ± SEM. Significance was set at P < 0.05.
To determine the localization of γ-cyto in mdx-TG skeletal muscle, cross-sections were stained with an affinity-purified γ-cyto actin antibody. Capillaries, blood vessels, nerves and immune cells stained brightly in mdx skeletal muscle (Figure 1B). In addition, the internal structure of mdx-TG muscle showed intense γ-cyto actin localization as previously described  for Actg1-TG mice (Figure 1B). Internal staining of γ-cyto actin was prominent in the majority of fibers. Staining of isolated mdx-TG myofibrils showed γ-cyto actin incorporation into the thin filament that was absent in mdx myofibrils (Figure 1C). Colocalization of γ-cyto actin with fast myosin heavy chain is consistent with reports that the human α-skeletal actin promoter used to drive transgene expression is preferentially active in fast-twitch muscle fibers [27, 36].
Isometric contractile properties of extensor digitorum longus muscles from four-month-old mdx and mdx-TG mice
mdx (n = 6)
mdx-TG (n = 6)
EDL mass (mg)
15.8 ± 0.8
13.7 ± 0.5
11.9 ± 0.2
12.2 ± 0.1
0.028 ± 0.001
0.024 ± 0.001
Passive stiffness (N/m)
19.6 ± 1.6
14.2 ± 0.9
Active stiffness (N/m)
714.8 ± 40.1
738.7 ± 98.3
59.6 ± 5.6
65.7 ± 8.1
17.9 ± 0.4
18.3 ± 0.4
21.4 ± 1.3
19.1 ± 0.7
Specific Pt (N/cm2)
2.1 ± 0.2
2.8 ± 0.4
264.0 ± 21.5
288.6 ± 39.1
7.15 ± 0.63
7.68 ± 0.95
10.9 ± 1.2
13.0 ± 2.4
Specific Po (N/cm2)
9.1 ± 0.6
12.2 ± 1.8
We are just beginning to understand the roles of cytoplasmic actin isoforms in skeletal muscle. Nonmuscle γ-cyto actin plays a vital role in muscle maintenance, as its ablation results in the accumulation of CNFs and hindlimb contractures . In dystrophic muscle, γ-cyto actin levels are elevated five- to tenfold compared to WT muscle, compensating for a weakened DGC and sarcolemma [23, 25]. When transgenically overexpressed 2, 000-fold (about 400 μM) in WT mice, γ-cyto actin incorporated into myofibrils but had no effect on isometric force generation, muscle fiber size or the incidence of CNFs . Similarly, we found that γ-cyto actin overexpression in dystrophic muscle also did not affect pathology, isometric force production or disease progression.
Our most interesting finding is that γ-cyto actin overexpression significantly decreased the immediate force loss caused by eccentric contractions in mdx muscle. This protection was apparent at two different ages of mice and was also associated with greater isometric force generation at the end of the eccentric contractions. Thus the protection afforded by γ-cyto actin overexpression was not due to weakened muscles, as isometric force production before the injury protocol was similar between mdx and mdx-TG mice and four-month-old mdx-TG mice actually generated greater eccentric force in seven of ten eccentric contractions compared to mdx mice. After eccentric contractions, EDL muscle mass was less in mdx-TG mice than in mdx mice, and there was a trend toward a greater passive stiffness increase in mdx mice than in mdx-TG mice, possibly suggesting reduced fluid accumulation as a consequence of muscle damage. These results differ from our findings in WT mice that Actg1-TG mice show a similar susceptibility to injury following eccentric contractions. It is possible that γ-cyto actin's protective effect in dystrophic muscle corrects a mechanism associated with force loss in dystrophic muscle that is not present in WT muscle.
The mechanism of force loss following eccentric contractions in dystrophic muscle is not understood but is likely different from that in healthy muscle. For example, in healthy muscle, excitation-contraction uncoupling has been demonstrated to account for about 75% of the deficit in muscle strength following eccentric contractions in vivo, with physical damage accounting for only approximately 25% . It has been postulated that extracellular calcium plays an important role in activating calpains to cause protein degradation. Furthermore, following excessive calcium entry into dystrophic muscle, the mitochondria act as a sink to buffer elevated calcium levels and consequently increase the production of reactive oxygen species [37–39]. Because the actin cytoskeleton can affect SOCE and interact with TRP channels , we hypothesized that γ-cyto actin's protective effect is due to its ability to attenuate extracellular calcium entry during eccentric contractions. Although mdx and mdx-TG mice appeared to have a slightly greater SOCE response than WT mice, we were unable to detect a difference between the three genotypes. In fact, the peak [Ca2+]i was not different between the genotypes, suggesting that any potential differences in SOCE were due to differences in basal [Ca2+]i. Our data differ from those reported by others who also used a calcium dye with other dystrophic models, such as mdx 5Cv and sarcoglycan δ-null mice [22, 34]. The mdx 5Cv mouse has fewer revertant fibers and an overall phenotype that is more severe than the mdx mouse . It is quite possible that calcium influx during store depletion is a phenotype seen in some mouse models of muscular dystrophy, but not in the mdx mouse.
Since we were not able to detect a difference in SOCE in immobilized FDB fibers, we performed more physiologically relevant experiments by modifying extracellular calcium or adding streptomycin to block stretch-activated channels to the muscle bath during ex vivo eccentric muscle contractions of the EDL muscle, similarly to previous work [12, 14, 15]. We found that removal of CaCl2 or the blockade of stretch-activated channels in the bathing media did not have an effect on immediate force loss during eccentric contractions. Our results are different from those previously published [12, 13, 15–17], probably because of the muscles studied (EDL vs FDB fiber or lumbrical muscle), differences in protocol (length change and rest time between contractions), length of treatment (acute vs chronic manipulation of genes) and calculation of force loss (eccentric vs isometric and timing of measurement).
The most commonly used muscle for the assessment of eccentric contraction-induced injury in mdx mice is the EDL , which we used in the current study. One previous study assessed the role of extracellular calcium and stretch-activated channel blockers during injurious muscle contractions in the EDL muscle , whereas all others have used FDB fibers or the lumbrical muscles [14–17]. It appears that isolated muscle fibers [14, 15] demonstrate a more robust response than intact muscles to stretch-activated channel blockers . Other studies supporting a role for extracellular calcium in force loss have been performed in the lumbrical muscle, using isometric contractions instead of eccentric contractions [16, 17].
Another difference between this and previous studies are the conditions of the eccentric injury protocol. Physiological length changes in muscle in a previous study were 20% or less . Researchers in other studies who have shown positive results by removing extracellular calcium or blocking stretch-activated receptors used length changes of 30% to 40% of Lo [12, 14, 15]. In the current study, we were able to use a physiological length change (Lo ± 5%) and still induce a significant amount of force loss. It is possible that a nonphysiological length change (more than 20% Lo) may stimulate a different mechanism responsible for force loss that is not apparent when the length change is relatively minor. The number of contractions and the rest time between contractions may also be important. The muscles in our study were rested for three minutes between contractions to prevent fatigue. Investigators in other studies have used rest periods ranging from 4 to 60 seconds between contractions. These short rest times can cause fatigue, which affects the amount of injury induced and complicates interpretation. With our three-minute rest period between contractions, we did not see any force loss during isometric or concentric contractions (data not shown).
Recent studies have suggested that TRP channels contribute to the dystrophic phenotype and contraction-induced injury [13, 21, 22]. Dystrophic mice harboring a dominant-negative TRPV2 or TRPC3 channel transgene show improved CK, fiber size variability, CNFs and susceptibility to contraction-induced injury [13, 21, 22]. In our study, acute blockage of stretch-activated channels with 2 mM streptomycin did not attenuate eccentric force loss. Mice that chronically overexpress a dominant-negative TRPV2 channel show improvement in CK, CNFs and contraction-induced injury [13, 21]. Perhaps chronic blockade of TRP channels leads to a healthier muscle, which indirectly results in protection from eccentric contractions.
It is also imperative to describe how force loss is calculated after eccentric contractions. Herein we report both eccentric force loss and isometric force loss. Although zero extracellular calcium or the addition of streptomycin had no effect on overall eccentric force loss, isometric force loss measured three minutes after the last eccentric contraction was significantly less in the zero calcium buffer but about threefold higher when treated with streptomycin. The reason for these disparate results may lie in the timing of the force measurements after injury. Whitehead et al.  did not report differences in eccentric force loss immediately after injury, but rather differences in isometric force loss 30 minutes after the last eccentric contraction when bathing EDL muscles in 0.2 mM streptomycin, which is similar to our finding. During that time, isometric force recovered from less than 10% of preinjury force to 34% to 44% of preinjury torque. We suggest that blocking stretch-activated channels enhances the recovery of muscle following eccentric contractions. Since some stretch-activated channels are nonselective for cations and zero extracellular calcium actually resulted in reduced isometric force (with or without EGTA), the effect of streptomycin on the recovery of force may be independent of calcium.
Our results provide new insight into the mechanism of contraction-induced injury in dystrophic muscle. First, γ-cyto actin overexpression can protect muscles from eccentric contraction-induced injury independently of any detectable improvement in dystrophic muscle pathology or disease phenotype. Second, γ-cyto actin overexpression appears to act through a mechanism independent of extracellular calcium or stretch-activated channels. Our data also suggest that dysfunction of stretch-activated cation channels is not involved in the immediate force loss following eccentric contractions in dystrophic muscle. Future studies will aim to determine the biochemical and biophysical properties of thin filaments, myofibrils and myofibers composed of different actin isoforms to gain understanding of how altered actin isoform composition translates into altered muscle function.
This work was supported by National Institutes of Health Training Program in Muscle Research grant AR007612, grants from the National Institutes of Health AR049899 (to JME), DA07304 (to SAT) and AG036827 (to DAL) and a grant from the National Science Foundation IOS0814549 (to SAT).
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