Expression of miR-133 in skeletal muscle.
The miR-133 family contains 3 highly homologous miRNAs: miR-133a-1, miR-133a-2, and miR-133b. miR-133a-1 and miR-133a-2 are identical and differ from miR-133b by 2 nucleotides at the 3′ terminus (18
). We have previously shown that miR-133a-1 and miR-133a-2 are expressed in cardiac and skeletal muscle, whereas miR-133b is skeletal muscle specific (18
). We determined the expression of miR-133 by Northern blot analysis in several skeletal muscles of different myofiber contents. Oxidative, type I (slow-twitch) myofibers are enriched in soleus muscle, and glycolytic type II (fast-twitch) myofibers are enriched in other muscle groups, such as gastrocnemius and plantaris (G/P), tibialis anterior (TA), and extensor digitorum longis (EDL) muscles. miR-133a was expressed at equivalent levels in all of these muscle groups (Figure A), indicative of its comparable levels in type I and type II myofibers. miR-133b was cotranscribed with miR-206 and was enriched in soleus muscle, which contains predominantly type I fibers (17
Expression of miR-133 in skeletal muscle.
We generated miR-133a–/–
(i.e., dKO) mice by interbreeding miR-133a-1+/–
mice, as described previously (18
), and confirmed the loss of miR-133a expression in dKO skeletal muscle by quantitative real-time RT-PCR (Figure B). The low level of miR-133 expression detected in dKO skeletal muscle represented the presence of miR-133b, which is detected by miR-133a probes. Based on results from real-time RT-PCR, we estimate that the relative abundance of miR-133a versus miR-133b in WT mice is about 15:1 in soleus and about 50:1 in G/P, EDL, and TA muscle, which confirms that miR-133b is less abundant than miR-133a in skeletal muscle and is enriched in soleus muscle.
Accumulation of centronuclear myofibers in dKO skeletal muscle.
dKO mice did not show apparent abnormalities in mobility. At 4 weeks of age, dKO muscles appeared normal by histological analysis and immunostaining for laminin and DAPI, and myofibers were comparable in size to those of WT muscle (Supplemental Figure 1, A–C; supplemental material available online with this article; doi:
). However, by 6 weeks of age, myofibers with centralized nuclei began to appear in dKO mice, and the percentage of myofibers with central nuclei in EDL, G/P and TA muscle increased progressively with age (Supplemental Figure 2A). By 12 weeks of age, nearly 60% of myofibers in TA muscle of dKO mice contained centralized nuclei (Figure , A–C). In contrast, dKO soleus muscle had relatively few centralized nuclei (Figure , A and C). These findings suggest that the phenotype of centrally located nuclei in dKO mice is specific to type II myofibers. In addition, at 12 weeks of age, dKO mice were significantly smaller in both body mass and mass of various muscle groups when normalized to tibia length (Supplemental Figure 2B). TA myofibers of dKO mice also had smaller diameters than normal at this age (Supplemental Figure 2C).
Centronuclear myofibers in dKO skeletal muscle.
As a further assessment of muscle abnormalities, we analyzed the distribution of mitochondria and sarcoplasmic reticulum (SR) by NADH-TR staining in dKO muscle fibers at 12 weeks. dKO fibers showed more oxidative enzyme activity in G/P, EDL, and TA muscles than did WT myofibers (Supplemental Figure 3A), which may reflect a shift from glycolytic to oxidative myofibers in these muscles. The oxidative enzyme activity within individual fibers was also unevenly distributed, and some myofibers showed radiating intermyofibrillary networks (Figure D). Ring-like fibers were also occasionally observed upon NADH-TR staining (Figure D). There was no significant difference in NADH-TR staining in soleus muscle between dKO and WT littermates (Supplemental Figure 3A). Interestingly, normal NADH-TR staining patterns were observed in 4-week-old dKO muscle when no centrally located nuclei were present (Supplemental Figure 3B).
Accumulation of centralized nuclei is usually indicative of muscle regeneration in response to disease or injury (21
). We therefore searched for signs of muscle damage and degeneration in dKO myofibers at 12 weeks of age. Monitoring sarcolemmal integrity by the uptake of Evans blue dye (EBD), which accumulates in damaged cells, showed very few dye-positive fibers (less than 4 per transverse section) (Figure E). We examined muscle from mdx mice, which develop muscular dystrophy, for comparison; these mice showed extensive EBD uptake (Figure E). We also measured serum levels of creatine kinase (CK) activity, indicative of sarcolemmal leakage, and observed only slightly elevated (2-fold) CK levels in dKO mice at 3 months (data not shown). In addition, dKO myofibers showed no signs of inflammation, fibrosis, or apoptosis (data not shown), which are characteristic of dystrophic muscle fibers. At 12 months of age, we did not observe worsening in myofiber morphologies or signs of inflammation, fibrosis, or cell death in dKO myofibers (Supplemental Figure 3C).
To assay for muscle regeneration, we analyzed expression of mRNAs encoding several myogenic markers of regeneration. Expression of Myog
(which encodes myogenin) was upregulated 7-fold in dKO TA muscle, but there was no change in the expression levels of other myogenic markers, such as Pax3
, and MyoD
(Figure F). Although there was a strong increase in both embryonic (Myh3
) and perinatal MHC (Mhy8
) mRNA levels in TA muscle by real-time RT-PCR (Figure F), embryonic MHC protein was rarely detected in dKO muscle fibers by immunohistochemistry (data not shown). These data indicate that there is only rare muscle regeneration in dKO mice, which is insufficient to account for the extensive centronuclear fibers observed in these mice. Thus, centronuclear myofibers in dKO mice without apparent necrosis, myofiber death, or significant regeneration are pathological characteristics reminiscent of human CNMs (1
T-tubule disorganization in muscle fibers of dKO mice.
In skeletal muscle, excitation-contraction coupling occurs at triads, which are composed of a transverse tubule (T-tubule) and a pair of terminal cisternae of the SR (24
). In Mtm1
-deficient mice, muscle fibers have a decreased number of triads and abnormal organization of T-tubules (3
). T-tubule disorganization has also been reported in human CNM patients (6
To assess whether T-tubule organization is affected in dKO muscle, we examined the expression of genes encoding components of T-tubules and SR that are important for excitation-contraction coupling, including the α1, β1, and γ1 subunits of the dihydropyridine receptor (DHPR) (encoded by Cacna1s
, and Cacng1
, respectively), ryanodine receptor 1 (Ryr1
), type 1 and 2 SERCA pumps (Atp2a1
), and calsequestrin 1 and 2 (Casq1
). At the mRNA level, expression of most of the genes was unchanged, except for a 2.5-fold increase in Cancng1
(Figure A). We also examined expression of RyR1, DHPRα, calsequestrin, and SERCA2 at the protein level and observed minimal changes (Supplemental Figure 4). In contrast, we observed a 35-fold increase in mRNA levels of Sln
, accompanied by a comparable increase in sarcolipin protein (Figure A and Supplemental Figure 4). Sarcolipin upregulation is a common feature in skeletal muscle myopathies (ref. 26
and M. Periasamy, unpublished observations), but the significance of this upregulation is unknown. Expression of phospholamban was slightly upregulated in dKO muscle, but the phosphorylated phospholamban was slightly decreased at the protein level (Supplemental Figure 4).
Disorganization of triads in TA muscle fibers in dKO mice.
We also analyzed the organization of triads by immunohistochemistry against DHPRα, a marker for T-tubules, and RyR1, a marker for terminal cisternae of SR. In transverse sections of WT myofibers, both T-tubules and terminal cisternae of SR displayed dot-like staining patterns distributed evenly along the myofibers (Figure B), which reflected the transverse orientations of triads relative to sarcomeres. In dKO myofibers, however, both T-tubules and SR showed aggregated staining, absence of staining in some regions, and irregular distribution within individual fibers (Figure B). In addition, in WT muscle, adjacent myofibers showed the same staining patterns. However, in dKO muscle, the adjacent myofibers often displayed different staining patterns (Figure B), suggestive of different orientations of triads in adjacent fibers. At 4 weeks of age, when dKO mice had not yet developed CNM, T-tubule structures were normal, as demonstrated by DHPRα staining (Supplemental Figure 3D).
We further analyzed the morphology of triads at the ultrastructural level by electron microscopy (Figure , C–J). In adult dKO TA muscle fibers, some T-tubules (stained dark by potassium ferricyanide) showed abnormal morphologies and longitudinal orientations aligned with the direction of myofibrils; these were rarely observed in WT muscle fibers (Figure , G–J). We also observed accumulation of electron-dense membranous structures along the myofibers and at triads in dKO myofibers (Figure , D–F). Overall, these findings indicate that miR-133a is important for the organization of T-tubules and triads and that its absence results in T-tubule disorganization.
Mitochondrial dysfunction in dKO skeletal muscle.
To determine whether lack of miR-133a alters mitochondrial function in skeletal muscle, mitochondria were isolated from red and white portions of the gastrocnemius muscle from dKO and WT mice. Immediately after isolation, mitochondrial respiration and fatty acid oxidation were assessed. Assessments of mitochondrial function include: (a) respiratory control ratio (RCR), the coupling between oxidative phosphorylation and ATP synthesis; (b) ADP-stimulated state 3 respiration, the respiratory rate during which the mitochondria are producing ATP; and (c) carbonylcyanide-p-trifluoromethoxyphenylhydrazone–stimulated (FCCP-stimulated) respiration, the maximal respiratory rate when oxidative phosphorylation is uncoupled from ATP synthesis. A reduction in any of these measures suggests mitochondrial dysfunction, which could be due to altered substrate handling, ATP synthase activity, or a dysfunction in respiratory chain components. Since we observed a reduction in all 3 parameters, dysfunction appears to be due to altered substrate handling or dysfunction in oxidative phosphorylation. The absence of miR-133a resulted in significant declines in RCR, ADP-stimulated state 3 respiration, and FCCP-stimulated maximal respiration in both red and white muscle, although the effects on FCCP-stimulated maximal respiration appeared to be more pronounced in red muscles (Figure A). In addition, total fatty acid oxidation was also significantly lower in mitochondria isolated from both red and white portions of gastrocnemius muscle from dKO animals (Figure B). There was also a reduction in citrate synthase in red quadricep muscle, but not in white quadricep muscle (Figure B). Collectively, these results demonstrate that the absence of miR-133a results in lower intrinsic mitochondrial function and fatty acid oxidation in both red and white skeletal muscle.
Mitochondrial dysfunction in dKO muscle.
miR-133a targets dynamin 2, a regulator of CNM.
To begin to explore the mechanistic basis of skeletal muscle abnormalities in dKO mice, we searched for targets of miR-133a with potential roles in CNM. Among the strongly predicted targets of miR-133a is Dnm2
mRNA, encoding a large GTPase implicated in endocytosis, membrane trafficking, and regulation of the actin and microtubule cytoskeletons (11
). Point mutations in the human DNM2
gene, thought to act in a dominant-negative manner, cause the autosomal-dominant form of CNM (7
). The 3′ UTR of Dnm2
mRNA contains an evolutionarily conserved miR-133a binding site (Figure A). miR-133a repressed a luciferase reporter gene linked to the 3′ UTR of Dnm2
mRNA, whereas a mutation in the predicted miR-133a binding site in the 3′ UTR prevented repression (Figure B), confirming Dnm2
mRNA as a target for miR-133a. Moreover, we observed a 2-fold increase in Dnm2
mRNA by quantitative real-time RT-PCR and an approximate 7-fold increase in dynamin 2 protein in TA muscle of dKO compared with WT mice by Western blot analysis (Figure , C and D). These results indicate that miR-133 represses dynamin 2 expression at both mRNA and protein levels.
miR-133a regulates Dnm2 expression in skeletal muscle.
Overexpression of dynamin 2 in skeletal muscle causes CNM in type II myofibers.
To examine whether elevated expression of dynamin 2, as observed in dKO myofibers, is sufficient to cause CNM, we generated transgenic mice in which dynamin 2 protein (with a myc-tag on the C terminus) was expressed under control of the muscle CK (MCK) promoter (referred to herein as MCK-DNM2 mice) (29
). Overexpression of dynamin 2 protein in skeletal muscle of transgenic mice was confirmed by Western blotting using antibodies against dynamin 2 as well as the myc epitope tag (Figure A). We obtained 2 MCK-DNM2 transgenic mouse lines, Tg1 and Tg2, which showed 3- and 6-fold overexpression of dynamin 2, respectively, compared with WT levels. At 7 weeks of age, both transgenic lines displayed accumulation of centronuclear myofibers (Figure B). Interestingly, Tg2 mice, which overexpressed dynamin 2 at a level similar to that of dKO mice, displayed age-dependent centronuclear myofibers in TA muscle comparable to those of dKO mice (Figure C).
Overexpression of Dnm2 in skeletal muscle causes CNM.
At 11 weeks of age, Tg2 mice displayed signs of muscle atrophy, with decreased muscle mass in both TA and G/P muscle (Supplemental Figure 5A). There was no difference in body mass between Tg2 and WT littermates (Supplemental Figure 5A). Histological analysis of TA muscle showed heterogeneous fiber sizes and the presence of centronuclear fibers in Tg2 mice (Figure D). The percentage of centronuclear myofibers in TA muscle of Tg2 mice was approximately 23% at this age (data not shown). NADH-TR staining revealed abnormal aggregation of oxidative enzymatic activity and radiating intermyofibrillary networks (Figure D). Abnormal organization of T-tubules was also observed in Tg2 TA muscle, as detected by immunohistochemistry against DHPRα (Supplemental Figure 5B).
Dynamin 2 protein was not significantly overexpressed in soleus muscle or heart of Tg2 mice (Supplemental Figure 5C), consistent with the preferential expression of the MCK promoter in type II myofibers (29
). Not surprisingly, therefore, we did not observe any abnormalities in soleus muscle or heart function in Tg2 mice (Supplemental Figure 5C and data not shown).
To assess muscle performance, we subjected mice to downhill treadmill running and analyzed running time and distance to exhaustion. At 10 weeks of age, Tg2 mice ran for a significantly shorter time than did WT mice (Figure E), indicative of muscle weakness. dKO mice showed a more dramatic decrease in running capacity (Figure E). However, the compromised cardiac function in dKO mice may also be a contributing factor to the reduction in exercise capacity.
Intracellular accumulation of dysferlin has been recently reported in human DNM2
-associated CNM patients, as well as in heterozygous mice carrying the R456W Dnm2
). We also analyzed localization of dysferlin in dKO muscle and Tg2 muscle. Interestingly, substantial accumulation of dysferlin inside the myofibers was observed in both dKO and Tg2 muscle fibers (Figure , A and B). Furthermore, at least some of the intracellular dysferlin was colocalized with dynamin 2 in dKO muscle fibers (Figure A).
Intracellular accumulation of dysferlin in dKO and MCK-DNM2 transgenic mouse myofibers.
These results demonstrate that elevated expression of Dnm2 in skeletal muscle causes CNM, predominantly in type II fibers, mimicking the dKO phenotype. We conclude that the CNM in dKO muscle can be explained, at least in part, by dysregulation of Dnm2.
dKO mice show increased type I myofibers in soleus muscle.
In addition to CNM, dKO mice displayed increased numbers of type I fibers in soleus muscle, which does not show CNM. We analyzed fiber type composition of soleus muscle from adult dKO mice by metachromatic ATPase staining and by immunohistochemistry against type I myosin heavy chain (MHC), shown by dark brown staining. Soleus muscle of WT mice was composed of about 43% type I fibers (Figure , A and B). Soleus muscle of dKO mice showed a 2-fold increase in the number of type I fibers (Figure , A and B).
Control of skeletal muscle fiber type by miR-133a.
Quantitative real-time RT-PCR analysis of the expression of transcripts encoding individual MHC isoforms revealed an increase in type I MHC (MHC-I) and decreases in type II MHCs (MHC-IIa, MHC-IIx/d, and MHC-IIb) in soleus muscle of dKO compared with WT mice (Figure C). We examined the protein composition of MHC isoforms in soleus, EDL, and TA muscle by silver staining of glycerol gels: 3 bands were present in protein extracts of soleus muscle isolated from WT mice, corresponding to MHC-IIa/IIx, MHC-IIb, and MHC-I proteins; 2 bands were present in protein extracts of TA and EDL muscles from WT mice, representing MHC-IIb and MHC-IIa/IIx (Figure C). Consistent with results from quantitative real-time RT-PCR, soleus muscle of dKO mice displayed an increase in MHC-I protein and a decrease in MHC-IIa/IIx proteins. MHC-IIb protein was not observed in dKO soleus muscle. Interestingly, there was an increase in the oxidative MHC-IIa/IIx protein and a decrease in the glycolytic MHC-IIb protein in TA and EDL muscles of dKO mice compared with WT mice, which indicates that these muscle groups also display a fiber type shift toward more oxidative (type IIa) fibers.
To determine whether loss of miR-133a affects the formation of type I fibers during fetal development, we examined MHC-I expression by immunohistochemistry at P1. There was no obvious difference in the number of MHC-I–positive myofibers in soleus or EDL muscles of dKO mice at P1 (Supplemental Figure 6A), which indicates that miR-133a does not influence embryonic development of type I myofibers. To determine when the fiber type switch takes place in dKO mice, we analyzed fiber type composition in both 2- and 4-week old mice by metachromatic ATPase staining. At both ages, the percentage of type I fibers in soleus was increased by almost 2-fold in dKO mice (Supplemental Figure 6B). We conclude that miR-133a does not influence specification of type I myofibers during embryonic development. Rather, miR-133a represses type I myofibers postnatally, such that the absence of miR-133a results in an increase in type I myofibers of adult mice.