Calcineurin signaling is essential for successful muscle regeneration. Although calcineurin inhibition compromises muscle repair, it is not known whether calcineurin activation can enhance muscle repair after injury. Tibialis anterior (TA) muscles from adult wild-type (WT) and transgenic mice overexpressing the constitutively active calcineurin-Aα transgene under the control of the mitochondrial creatine kinase promoter (MCK-CnAα*) were injected with the myotoxic snake venom Notexin to destroy all muscle fibers. The TA muscle of the contralateral limb served as the uninjured control. Muscle structure was assessed at 5 and 9 days postinjury, and muscle function was tested in situ at 9 days postinjury. Calcineurin stimulation enhanced muscle regeneration and altered levels of myoregulatory factors (MRFs). Recovery of myofiber size and force-producing capacity was hastened in injured muscles of MCK-CnAα* mice compared with control. Myogenin levels were greater 5 days postinjury and myocyte enhancer factor 2a (MEF2a) expression was greater 9 days postinjury in muscles of MCK-CnAα* mice compared with WT mice. Higher MEF2a expression in regenerating muscles of MCK-CnAα* mice 9 days postinjury may be related to an increase of slow fiber genes. Calcineurin activation in uninjured and injured TA muscles slowed muscle contractile properties, reduced fatigability, and enhanced force recovery after 4 min of intermittent maximal stimulation. Therefore, calcineurin activation can confer structural and functional benefits to regenerating skeletal muscles, which may be mediated in part by differential expression of MRFs.
- myotoxic injury
- skeletal muscle
- muscle function
calcineurin is a serine/threonine phosphatase enzyme that regulates transcription by sensing changes in intracellular calcium-calmodulin concentrations (32). The calcineurin signal transduction pathway is important in skeletal muscle biology, because it can regulate the slow oxidative phenotype, regeneration, and myofiber growth (11, 16, 29). In animal models, pharmacological inhibition of calcineurin impairs skeletal muscle regeneration after injury (1, 38), and in regenerating muscles of young mdx dystrophic mice, it can severely compromise muscle structure and function (44). The effect of calcineurin stimulation on muscle regeneration efficacy is unknown.
Calcineurin controls satellite cell differentiation and myofiber growth and maturation. Activated calcineurin promotes the transcription and activation of myoregulatory factors (MRFs), myocyte enhancer factor 2 (MEF2), myogenin, and MyoD, either directly or in conjunction with nuclear factor of activated T cells (NFAT) (6, 11, 12, 38). The NFATc1, -c2, and -c3 isoforms are expressed in skeletal muscle (15) and have been observed in the cytoplasm of muscle cells at all stages of development (1). Specificity may exist among the NFAT isoforms in gene regulation, since each isoform has been shown to translocate to the nucleus at specific stages of muscle differentiation (1). Myogenin, MyoD, and MEF2 stimulate the differentiation of satellite cells and the maturation of myoblasts into myotubes (30). MEF2a also regulates the expression of the developmental myosin heavy chain (MyHC) isoform in newly regenerated fibers (35) and slow fiber-type-specific genes (49). In maturing muscle fibers, calcineurin can alter the expression of adult type I and IIa MyHC isoforms (2, 48).
Calcineurin has been implicated in regulating myofiber growth, and a positive relationship between calcineurin activation and muscle hypertrophy has been demonstrated (25). In myocyte cultures, calcineurin activation enhanced myotube formation (5). Cells expressing the activated calcineurin-Aα (CnAα*) transgene were larger and contained more myonuclei, and MyHC protein and the hypertrophic response was associated with increased NFATc3 nuclear translocation (5). In vitro, IGF-I has been shown to activate calcineurin and its downstream transcription factors NFATc1 and GATA to stimulate myocyte hypertrophy (28, 42). Pharmacological inhibition of calcineurin prevented hypertrophy following functional overload of mouse plantaris muscle (9, 10). However, in vitro, the contribution of calcineurin signaling to muscle hypertrophy is influenced by experimental design (41). In experiments where the hypertrophic response depends on myoblast maturation and fusion (42) or the addition of myonuclei (26), the involvement of the calcineurin signal transduction pathway appears more definite. During development, myoblasts fuse to form nascent myotubes with a limited number of myonuclei. Further growth of these nascent myofibers depends on the fusion of additional myonuclei (16), and this can be controlled by calcineurin and NFATc2 (19). Activation and nuclear translocation of NFATc2 in nascent myotubes stimulates IL-4 transcription and synthesis (18). Secreted IL-4 is thought to bind to its receptor on myoblasts to initiate fusion of myoblasts to nascent myotubes (18). The physiological significance of this pathway has been demonstrated in cell cultures and NFATc2−/− knockout mice (16, 17, 19). In vivo, NFATc3 may regulate early events in primary myogenesis, given that muscles from NFATc3−/− knockout mice contain fewer myofibers than those from WT mice; however, mean fiber size is normal in these mice (21).
It is important to note that the effect of calcineurin activation on growth and hypertrophy is strongly influenced by muscle phenotype. Compared with wild-type (WT) mice, slow-twitch muscles (e.g., soleus) of mice overexpressing an activated form of calcineurin (MCK-CnAα*) have greater mass, and their mean fiber cross-sectional area is greater (47). However, fast-twitch muscles (e.g., tibialis anterior and medial gastrocnemius) of MCK-CnAα* mice have a smaller mass and fiber size compared with muscles from WT mice, and there is no change in mass or fiber size in muscles with an intermediate phenotype (e.g., plantaris) (47). Furthermore, genetic inhibition of calcineurin by muscle-specific overexpression of modulatory calcineurin-interacting protein 1 (MCIP1) reduced fiber size in slow-twitch soleus muscles by 30% but had no effect on fiber size in any other muscle type (31).
The aim of this study was to investigate whether expression of the constitutively active CnAα* transgene would improve muscle structure and function in regenerating muscles following myotoxic injury. It was hypothesized that increased calcineurin activity would hasten muscle regeneration by stimulating satellite cell differentiation and fusion of differentiated satellite cells to nascent myofibers, accelerating recovery of myofiber size. Furthermore, it was hypothesized that calcineurin activation would stimulate the maturation of newly regenerated myofibers, thus enhancing functional recovery.
All experiments were approved by the Animal Experimentation Ethics Committee of The University of Melbourne. The MCK-CnAα* mice express an active from of calcineurin-Aα (CnAα*). The CnAα* transgene is driven by the mitochondrial creatine kinase (MCK) promoter, which is induced during myogenic differentiation, and its activity is maximal in committed myotubes (43). The transgenic mice were developed at the Department of Molecular Biology at the University of Texas Southwestern Medical Center at Dallas (29). To generate the animals used in this study, the MCK-CnAα* mice, which have a B6C3F1 background, were mated with B6C3F1 WT mice. Half of the offspring expressed the CnAα* transgene (MCK-CnAα*), and the others were used as WT controls.
Mice that expressed the CnAα* transgene were identified through PCR screening of DNA extracted from tail tissue (29). Mice were maintained on a 12:12-h light-dark cycle, with standard mouse chow and water provided ad libitum. Assessments of tibialis anterior (TA) muscle function and regeneration were conducted on MCK-CnAα* and WT mice at 8.5 ± 0.8 mo of age.
Muscle regeneration was assessed at 5 and 9 days postinjury. In the 5-day-postinjury group, the right TA was injured and the left TA muscle served as the uninjured control for assessment of muscle structure and function. In the 9-day-postinjury group, the left TA muscle was injured and used for muscle function testing and the right TA muscle served as the uninjured control. Histological and immunohistochemical analyses were performed on the injured and uninjured muscles at 5 and 9 days postinjury. Schertzer and Lynch (40) showed previously that after Notexin damage, muscle function can be measured reliably at 7 days postinjury. Before this time, the structural integrity of the degenerating-regenerating muscle does not permit consistent and reliable assessments of force-producing capacity. Mice were anesthetized with 100 mg/kg ketamine (Parnell Laboratories, Alexandria, NSW, Australia) and 10 mg/kg xylazine (Troy Laboratories, Smithfield, NSW, Australia) such that they were unresponsive to tactile stimuli. The appropriate (right or left) TA muscle was surgically exposed and injected to maximal holding capacity with the myotoxin Notexin (1.0 μg/ml, ∼50 μl; Latoxan, Valence, France) through two injections in the proximal and distal regions of the muscle with a 30-gauge needle. The incision was closed with Michel clips (Aesculap, Tuttlingen, Germany), and the mice were kept warm on a heat pad and monitored closely until they regained consciousness. Injecting a muscle with Notexin destroys all mature muscle fibers but leaves satellite cells intact to facilitate muscle regeneration (14). The contralateral uninjured TA muscle was not injected with a saline vehicle, since previous studies have shown that injecting muscles with saline does not affect muscle function or structure (13, 20, 36).
Assessment of muscle function in situ.
Details regarding the in situ TA muscle preparation and measurement of contractile properties have been described previously (7, 40). Briefly, the anesthetized mouse was placed on a heated (37°C) platform, the left foot of the mouse was secured to the platform, and the left knee was secured by pushing a sharpened pin attached to a clamp through the knee joint. The distal portion of the muscle and its tendon were dissected free and maintained at 37°C by incubating the exposed muscle in warmed mineral oil. The tendon was tied with 4-0 braided surgical silk to the lever arm of a dual-mode servomotor/force transducer (305-LR; Aurora Scientific, Aurora, ON, Canada). The muscle was stimulated maximally by directly applying supramaximal square wave pulses of 10 V, 300 ms in duration, to the exposed nerve (40). Optimum muscle length (Lo) was determined from micromanipulations of muscle length and a series of isometric twitch contractions. Maximum isometric tetanic force (Po) production was determined from the plateau of the frequency-force relationship. To assess fatigability, Po was determined once every 5 s for 4 min, and the muscle was then rested (no stimulation). Recovery of Po was determined at 1, 2, and 3 min after the fatigue protocol. Optimal fiber length (Lf) was determined by multiplying Lo by the TA Lf/Lo ratio of 0.6 (7).
After the final evaluation of Po, the TA muscle was carefully dissected from the mouse, trimmed of tendon and suture, and weighed on an analytical balance. The contralateral TA muscle was also excised. Both muscles were mounted separately in embedding medium, frozen in thawing isopentane, and stored at −80°C for later histological and immunohistochemical analyses. The mice were killed by cardiac excision while still anesthetized deeply. Muscle cross-sectional area was calculated by dividing muscle mass by the product of Lf and 1.06 mg/mm3, the density of mammalian skeletal muscle. Force per cross-sectional area, or specific force (sPo, kN/m2), was determined by dividing Po by muscle cross-sectional area (24).
Histology and immunohistochemistry.
Four 5-μm-thick sections were cut from the midbelly region of each muscle on a cryostat at −20°C. The sections were air-dried overnight and stored at −80°C. One section was stained with hematoxylin and eosin (H&E) for determination of fiber cross-sectional area, muscle degeneration, and the proportion of centrally nucleated fibers, and the others were used for immunohistochemical analysis. Colored, digitized images of H&E-stained sections were captured with a digital camera (Spot, v2.2; Diagnostic Instruments, Sterling Heights, MI) mounted to an upright microscope (BX-51; Olympus, Tokyo, Japan) at ×200 magnification for mean fiber cross-sectional analysis and at ×100 magnification for assessment of TA muscle degeneration and regeneration and to determine the number of fibers per area (mm2) of tissue. Images were analyzed using an image analysis system (AIS, v6.0; Imaging Research, St. Catherines, ON, Canada). Mean fiber cross-sectional area was calculated by analyzing ∼150 fibers per section. To quantitate regeneration efficacy in TA muscles at 9 days postinjury, areas of degeneration were identified and manually outlined using the image analysis program and expressed as a percentage of total section area examined; ∼3 mm2 of muscle section were examined per sample.
Immunohistochemical techniques were used to assess markers of muscle regeneration in WT and MCK-CnAα* mice. Muscle sections were reacted with antibodies against myogenin (no. sc-576, diluted 1:60; Santa Cruz Biotechnology), MEF2a (no. sc-313, diluted 1:60; Santa Cruz Biotechnology), and the developmental MyHC isoform (no. NLC-MHCd, diluted 1:20; Novocastra) as described previously (45). The number of MEF2a- and myogenin-positive nuclei in each section was counted and expressed as the number of positive nuclei per area (mm2) of muscle. The number of muscle fibers that reacted with the anti-developmental MyHC antibody was counted and expressed as a proportion of the total number of muscle fibers (616 ± 50 fibers examined per section); their cross-sectional area was also determined. We reacted cross sections from uninjured muscles and from injured muscles at 5 and 9 days postinjury with the developmental MyHC antibody but did not present the 5-day-post injury data for two reasons. First, at 5 days postinjury, almost all of the fibers expressed the developmental MyHC, which made it difficult to discern the effect of calcineurin-Aα activation on developmental MyHC expression. These fibers were (completely) stained red, unlike those from the 9-day-postinjury samples, in which some fibers had only partial staining for developmental MyHC. Second, the MyHC antibody (no. NLC-313; Novacastra) is a mouse monoclonal antibody, and the secondary rabbit anti-mouse antibody (no. E0464; Dako, Botany, Australia) can cross-react with endogenous mouse immunoglobulins (IgGs). Visualization of developmental MyHC staining on mouse muscle cross sections was achieved using a peroxidase detection system (Vectastain ABC kit, no. PK-6100; Vector Laboratories) and the AEC+ substrate chromogen (no. K3468; Dako) for color development. To control for nonspecific staining due to secondary antibody cross-reactivity, one negative control section from WT uninjured muscles incubated with the secondary antibody and the Vectastain kit and another negative control incubated with the Vectastain kit only were included with each staining run. In uninjured muscle sections from WT mice, nonspecific staining was minimal and confined to the interstitium and not the myofiber. At 5 days postinjury, there was still a considerable inflammatory response, with infiltration of neutrophils and macrophages to remove necrotic myofibers and promote regeneration. This was associated with increased IgG levels in the regenerating muscle and greater nonspecific staining in the interstitium, making it difficult to discern myoblast and regenerating fibers from inflammatory cells and cellular debris exposed to mouse IgGs. All sections for immunohistochemical analysis were counterstained with hematoxylin, which stained nuclei blue.
Values are means ± SE. All statistical analyses were performed using a statistics software package (Minitab, State College, PA). ANOVA or two-sample t-tests were used to assess differences between uninjured and 5- or 9-day-postinjury TA muscles from MCK-CnAα* and WT mice. Tukey's post hoc test was used to locate pairwise significant differences, where appropriate. For all comparisons, P < 0.05 was considered significant.
Body and muscle mass of MCK-CnAα* mice.
Body mass was similar in MCK-CnAα* and WT mice. TA muscles from MCK-CnAα* mice had a lower mass than those from WT mice (P < 0.05), and the muscle mass to body mass ratio (MM/BM) was lower (P < 0.05). Nine days after myotoxic injury, muscle mass from both strains was ∼30% less than that of uninjured muscles and of injured muscles at 5 days postinjury (P < 0.05; Table 1).
Muscle function is improved after injury in MCK-CnAα* mice.
Twitch force (Pt) tended to be greater (∼5%) in uninjured muscles from MCK-CnAα* than WT mice (P = 0.055). Nine days postinjury, Pt was ∼35% that of uninjured Pt (P < 0.05). Po was ∼20% lower in uninjured muscles from MCK-CnAα* than in those from WT mice (P < 0.05; main effect). Nine days post injury, Po was reduced by 71% compared with uninjured Po in MCK-CnAα* and WT mice (P < 0.05). Normalized force (sPo) of uninjured muscles from MCK-CnAα* mice was ∼15% less than that of WT mice (P < 0.05). However, 9 days after injury, sPo of muscles from MCK-CnAα* and WT mice was similar: 44 and 35% of uninjured sPo, respectively (Table 1). Time to peak tension (TPT) and half-relaxation time (½RT) were prolonged in injured muscles (P < 0.05; main effect) and in muscles expressing the CnAα* transgene compared with WT mice (P < 0.05; Table 1).
Muscle fatigue is decreased in muscles of MCK-CnAα* mice.
TA muscles of MCK-CnAα* mice fatigued less during 4 min of intermittent maximal stimulation and recovered their force-generating capacity better than muscles of WT mice (P < 0.05). At 2, 3, and 4 min into the fatigue protocol, relative Po was higher in muscles from MCK-CnAα* than in muscles from WT mice (P < 0.05). At 1, 2, and 3 min into recovery, Po of muscles from MCK-CnAα* mice had returned to initial values (time = 0 min of fatigue), whereas Po of muscles from WT mice never fully recovered (P < 0.05). Uninjured muscles tended to fatigue more than injured/regenerating muscles during the repeated stimulation protocol, but force recovery following the 4-min fatigue protocol tended to be superior than injured muscles (not significant; Fig. 1A). It should be noted that injured muscles produced less force than uninjured muscles during the fatigue protocol and recovery (Fig. 1B).
Muscle degeneration is decreased in MCK-CnAα* mice after injury.
In uninjured muscles of MCK-CnAα* mice, mean fiber size was ∼30% smaller than in WT mice (P < 0.05). Five days postinjury, mean fiber size was 23% of control values in muscles from WT and MCK-CnAα* mice. At 9 days postinjury, muscles from MCK-CnAα* mice recovered their mean (uninjured) fiber size to a greater extent than muscles from WT mice (49 vs. 39%) such that the difference in fiber size between MCK-CnAα* and WT mice was ∼16% and no longer statistically significant (P > 0.05; Table 2). Muscle degeneration, defined as areas of connective tissue, fat, and mononuclear cell infiltration, was assessed at 9 days postinjury. In muscles of MCK-CnAα* mice, degeneration occupied 38% of the cross section analyzed, whereas in WT mice it was 51% (P < 0.05; Fig. 2, A and D).
Expression of the CnAα* transgene appeared to increase myofiber number in uninjured and regenerating muscles (P < 0.05), with the number of fibers per area (mm2) of tissue being 42% greater in uninjured muscles and 33% greater in muscles of MCK-CnAα* mice compared with WT mice at 9 days postinjury. In both MCK-CnAα* and WT mice, the number of muscle fibers was higher in injured than in uninjured muscle (P < 0.05; Fig. 2C).
Expression of myogenic markers is increased in MCK-CnAα* mice during early regeneration.
In WT and MCK-CnAα* mice, the number of myogenin- and MEF2a-positive nuclei was significantly higher in injured than in uninjured TA muscles (P < 0.05; Figs. 3 and 4). Five days postinjury, myogenin expression was higher in muscles of MCK-CnAα* than WT mice (P < 0.05). Between 5 and 9 days postinjury, myogenin expression was decreased approximately fourfold in muscles of MCK-CnAα* and approximately twofold in muscles of WT mice such that by 9 days postinjury, myogenin expression was similar in both strains (Fig. 3). Similarly, the number of myogenin-positive nuclei per muscle fiber (as assessed in cross section) was lower in the muscles of MCK-CnAα* compared with WT mice at 9 days postinjury (data not shown). At 9 days postinjury, the number of MEF2a-positive nuclei was higher in muscles of MCK-CnAα* than in WT mice (P < 0.05; strain × injury interaction; Fig. 4), but not when the number of MEF2a-positive nuclei was expressed per myofiber (data not shown). MEF2a expression was highest 9 days postinjury in MCK-CnAα* mice (P < 0.05) and 5 days postinjury in WT mice (P = 0.09).
At 9 days postinjury, the proportion of centrally nucleated fibers and the number of central nuclei per fiber were similar in muscles from MCK-CnAα* and WT mice (Fig. 2B). The proportion of fibers expressing the developmental MyHC isoform was higher in injured than in uninjured muscles (P < 0.05) and similar in regenerating muscles from MCK-CnAα* and WT mice (Fig. 5).
Calcineurin activation was associated with the upregulation of specific MRFs in regenerating skeletal muscles. Five days post myotoxic injury, myogenin expression was greater and 9 days postinjury, MEF2a levels were greater in muscles from MCK-CnAα* mice than in WT mice. The alterations in MRFs were associated with the enhanced regeneration and maturation of myofibers, since at 9 days postinjury, recovery of fiber size, overall muscle architecture, and relative normalized force-producing capacity were superior in muscles from MCK-CnAα* mice compared with WT mice.
Calcineurin activation causes a shift to a slower muscle phenotype (22, 29, 47) and an increased expression of type IIa and I MyHC isoforms and the slow sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA) 2a isoform in skeletal muscle (3, 4). These effects were associated with concomitant changes in muscle function (46). Similarly, in the present study, twitch contraction time (TPT) and relaxation time (½RT) were prolonged in TA muscles of MCK-CnAα* compared with WT mice. Consistent with a slower phenotype, muscles from MCK-CnAα* mice had an altered twitch/tetanus ratio, fatigued less during repeated contractile activity, and exhibited an improved force recovery. The shift to a slower phenotype reduced the force-producing capacity, as evident from the ∼20% decrease in Po and ∼15% decrease in sPo in muscles of MCK-CnAα* mice. These changes were attributed to the ∼15% decrease in muscle mass and ∼30% decrease in fiber size. In TA muscles of MCK-CnAα* mice, the decrease in fiber size was twofold greater than the decrease in muscle mass. This can be accounted for by the 42% increase in myofiber number. In regenerating muscles of MCK-CnAα* mice, myofiber number was increased by 28%. We previously observed a twofold increase in myofiber number in muscles of CnAα* mdx (dystrophic) mice (46). Others have reported an ∼40% decrease in myofiber number in soleus muscles from CnAα knockout mice (34). Changes in myofiber number in response to genetic manipulation of calcineurin may be secondary to changes in fiber size (34), or they may be mediated by greater NFATc3 activation (21), since NFATc3 can regulate myofiber number during primary myogenesis (21).
Calcineurin activation may enhance the maturation of regenerating muscle fibers after myotoxic injury through increased MRF expression. In regenerating muscles from MCK-CnAα* mice, a greater proportion of the muscle cross-sectional area was occupied by viable muscle fibers and less was occupied by noncontractile tissue (inflammatory cell, connective and adipose tissue). Furthermore, the relative recovery of regenerating fiber size was superior. These factors likely contributed to the enhanced recovery of relative sPo in muscles from MCK-CnAα* mice: 44% of uninjured values compared with 35% in WT mice.
Expression of myogenin and MEF2a is regulated, at least in part, by the calcineurin signal transduction pathway (12, 49, 50). Calcineurin activation increased myogenin expression in regenerating muscles 5 days postinjury. This finding was surprising given that the CnAα* transgene is driven by the MCK promoter, which is most active in fused myotubes (43). At 9 days postinjury, MEF2a levels were greater in MCK-CnAα* than in WT mice. In uninjured muscles of adult MCK-CnAα* and WT mice, myogenin and MEF2a levels were similar. Others have also observed similar levels of MEF2 activation in muscles from CnA-deleted mice and WT mice (33), suggesting that in adult mice, fiber type proportions are set and genetic manipulation of calcineurin does not influence downstream MRF expression. Between 5 and 9 days postinjury, myogenin expression was downregulated in regenerating muscles, and this reduction was twofold greater in MCK-CnAα* than in WT mice. Within that time frame, MEF2a expression was downregulated in regenerating muscles of WT mice but upregulated in regenerating muscles of MCK-CnAα* mice. Despite greater MEF2a levels at 9 days postinjury in muscles from MCK-CnAα* mice, there was no accompanying increase in the levels of developmental MyHC. The more rapid downregulation of myogenin expression and higher levels of MEF2a with no concomitant increase in the levels of developmental MyHC expression suggests that regeneration was accelerated in MCK-CnAα* compared with WT mice and that MEF2a may have been regulating the expression of slow fiber-specific genes, rather than developmental genes.
The cellular mechanisms responsible for the superior recovery of muscle morphology in regenerating muscles expressing the CnAα* transgene remains to be elucidated. At 9 days postinjury, the proportion of centrally nucleated myofibers and the number of central nuclei per myofiber were similar in MCK-CnAα* and WT mice, suggesting that in fast muscles, calcineurin activation may not increase the number of myoblasts fusing to regenerating myofibers. This does not preclude the possibility that secondary fusion occurs sooner in muscles expressing the CnAα* transgene. A limitation of our “proportion of centrally nucleated fibers” analysis is that it does not account for myofiber number. Although the proportion of centrally nucleated fibers was similar in regenerating TA muscles of MCK-CnAα* and WT mice, total myofiber number was ∼28% greater, and the number of centrally nucleated fibers (i.e., the fusion index) was ∼24% greater, in muscles of MCK-CnAα* compared with WT mice. This raises the issue of whether the proportion of centrally nucleated fibers or the fusion index is a more valid measure for assessing the effect of calcineurin activation on myoblast fusion. Calcineurin activation could enhance secondary myoblast fusion in regenerating muscles through the NFATc2 and IL-4 pathway (17, 18).
We did not detect any deleterious side effects of calcineurin activation in skeletal muscle. In cardiac muscle, depending on the experimental model, calcineurin activation can either inhibit or enhance apoptosis (27). Cardiomyopathy has been observed in transgenic mice expressing a constitutively active calcineurin transgene (23). Some of these effects have been attributed to mitochondrial dysfunction and elevated superoxide production (39). Mitochondrial biogenesis is a crucial regulatory event during muscle regeneration. In regenerating rat muscles, the most prominent increase in mitochondrial respiratory capacity occurs between 5 and 9 days post myotoxic injury, presumably to coincide with the increased metabolic cost of regeneration (8). In our study, differences in regeneration efficacy between MCK-CnAα* and WT mice became apparent between 5 and 9 days post injury, making it unlikely that regeneration was compromised by mitochondrial dysfunction or increased reactive oxygen species production. The superior fatigue resistance and increased fatty oxidation capacity of muscles from MCK-CnAα* compared with WT mice (37) offer further indirect support for normal mitochondrial function with calcineurin activation in skeletal muscle. Nonetheless, the possibility that calcineurin activation could increase oxidative stress in regenerating skeletal muscles is interesting and worthy of further examination.
In conclusion, activation of the calcineurin signal transduction pathway enhanced regeneration in muscles after myotoxic injury and promoted recovery of force-producing capacity. The cellular mechanisms responsible for the improvement in muscle structure and the enhanced recovery of fiber size have yet to be elucidated and may involve alterations in the timing of the MRFs. In addition, any potential side effects of calcineurin activation need to be assessed carefully (39). Our findings are consistent with the hypothesis that with calcineurin activation, maturation of regenerating myotubes is more effective.
This work was supported by research grants from the National Health & Medical Research Council of Australia (project numbers 350439 and 454561).
Present address for N. Stupka: Institute of Biotechnology (BioDeakin), Deakin University, Geelong, Victoria 3217, Australia (e-mail:).
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