Mice lacking microRNA 133a develop dynamin 2–dependent centronuclear myopathy (original) (raw)
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 1A), 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. (A) Northern blot analysis of miR-133a in adult WT mouse tissues. The blot was stripped and reprobed with 32P-labeled U6 probe as a loading control. Sol, soleus. (B) Expression of miR-133 in skeletal muscle, detected by real-time RT-PCR and expressed relative to U6.
We generated miR-133a–/– (i.e., dKO) mice by interbreeding miR-133a-1+/–;miR-133a-2+/– mice, as described previously (18), and confirmed the loss of miR-133a expression in dKO skeletal muscle by quantitative real-time RT-PCR (Figure 1B). 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:10.1172/JCI46267DS1). 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 2, A–C). In contrast, dKO soleus muscle had relatively few centralized nuclei (Figure 2, 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. (A) H&E staining of soleus, EDL, G/P, and TA muscles of WT and dKO mice at 12 weeks of age. Scale bars: 40 μm. (B) Immunostaining of TA muscle against laminin. Nuclei are stained with DAPI. dKO TA muscle showed central nuclei. Scale bars: 40 μm. (C) Percentage of centronuclear myofibers in 4 WT mice and 10 dKO mice at 12 weeks of age. For each mouse, more than 500 myofibers were counted for TA and G/P muscles and more than 300 myofibers were counted for soleus and EDL muscles. (D) NADH-TR staining of dKO TA muscle revealed abnormal distribution, radiating intermyofibrillary network (arrows), and ring-like fibers (asterisks). Scale bars: 20 μm. (E) EBD uptake of TA muscles of WT, dKO, and mdx mice. Immunostaining with laminin (green) is shown; EBD is detected as a red signal under fluorescence microscopy. Scale bars: 100 μm. (F) Expression of myogenic genes and of embryonic MHC (Myh3) and perinatal MHC (Myh8) in WT and dKO TA muscle, determined by real-time RT-PCR. n = 3 (WT and dKO).
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 2D). Ring-like fibers were also occasionally observed upon NADH-TR staining (Figure 2D). 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 (21–23). 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 2E). We examined muscle from mdx mice, which develop muscular dystrophy, for comparison; these mice showed extensive EBD uptake (Figure 2E). 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, Pax7, and MyoD (Figure 2F). 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 2F), 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, 2).
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, 25).
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, Cacnb1, and Cacng1, respectively), ryanodine receptor 1 (Ryr1), type 1 and 2 SERCA pumps (Atp2a1 and Atp2a2), and calsequestrin 1 and 2 (Casq1 and Casq2). At the mRNA level, expression of most of the genes was unchanged, except for a 2.5-fold increase in Cancng1 (Figure 3A). 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 3A 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. (A) Expression of mRNA transcripts encoding components of T-tubules and SR was determined by real-time RT-PCR in TA muscles of 12-week-old mice. n = 3 (WT and dKO). (B) Immunostaining of T-tubules and SR in transverse sections of TA muscle from WT and dKO mice at 12 weeks of age. T-tubules were detected by anti-DHPRα, and terminal cisternae of the SR were detected by anti-RyR1. Nuclei were detected by DAPI, and the myofiber perimeter was stained by anti-laminin. Images of multiple levels of the sections were taken and reconstructed to create the 3D effect. Scale bars: 30 μm. (C–J) Electron micrographs of WT and dKO muscle. dKO TA muscle showed accumulation of electron-dense structures (D–F) that were absent in WT TA muscle (C). dKO muscle (H and J) displayed T-tubules (arrows) in abnormal orientations compared with WT muscle (G and I). Scale bars: 2 μm (C and D); 0.5 μm (E–H); 0.2 μm (I and J).
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 3B), 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 3B). 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 3B), 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 3, 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 3, G–J). We also observed accumulation of electron-dense membranous structures along the myofibers and at triads in dKO myofibers (Figure 3, 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 4A). 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 4B). There was also a reduction in citrate synthase in red quadricep muscle, but not in white quadricep muscle (Figure 4B). 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. (A) Mitochondria were isolated from red and white gastrocnemius muscle, and oxygen consumption rate (OCR) was measured for RCR, ADP-stimulated state 3 respiration (ADP), and FCCP-stimulated respiration (FCCP). n = 6 (WT and dKO). *P < 0.05 vs. WT. (B) Fatty acid oxidation was measured in isolated mitochondria from red and white gastrocnemius muscle. Citrate synthase enzyme activity was measured in isolated mitochondria from red and white quadriceps muscle. n = 6 (WT and dKO). *P < 0.05 vs. WT.
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, 8, 27, 28). The 3′ UTR of Dnm2 mRNA contains an evolutionarily conserved miR-133a binding site (Figure 5A). 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 5B), 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 5, 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. (A) Position of miR-133a target site in Dnm2 3′ UTR and sequence alignment of miR-133a and the Dnm2 3′ UTR from various species are shown. Conserved miR-133a binding sites in Dnm2 3′ UTR are shown in red. Mutations in Dnm2 3′ UTR were introduced to disrupt base-pairing with miR-133a seed sequences (blue). (B) Luciferase reporter constructs containing WT and mutant Dnm2 3′ UTR sequences were cotransfected into COS-1 cells with a plasmid expressing miR-133a. 48 hours after transfection, luciferase activity was measured and normalized to β-galactosidase activity. (C) Real-time RT-PCR showing expression of Dnm2 mRNA in WT and dKO TA muscle. n = 3 (WT and dKO). (D) Western blot showing expression of dynamin 2 protein in TA muscle of WT and dKO mice. n = 2 (WT and dKO). The blot was stripped and reprobed with an antibody against α-actin as a loading control. Quantification of dynamin 2 protein, determined by densitometry and normalized to α-actin, is also shown.
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, 30). 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 6A). 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 6B). 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 6C).
Overexpression of Dnm2 in skeletal muscle causes CNM. (A) Western blot analysis of TA muscle from WT and MCK-DNM2 transgenic mouse lines Tg1 and Tg2 using anti–dynamin 2 and anti-myc to show overexpression of transgene. Anti-tubulin was used as a loading control. Protein quantification, determined by densitometry, is also shown. (B) Transverse sections of TA muscles of WT, Tg1, and Tg2 mice at 6 weeks of age were stained with wheat germ agglutinin (WGA) and DAPI to show central nuclei (arrows) in transgenic mice. Scale bars: 100 μm. (C) Percentage of centronuclear myofibers in TA muscle of transgenic mice at 7 weeks of age. (D) Histological analysis of TA and soleus muscles of WT and Tg2 mice at 11 weeks of age. TA muscle sections were stained with H&E, with anti-laminin and DAPI to show central nuclei, and with NADH-TR to reveal abnormal distribution and radiating intermyofibrillary network (arrows). Scale bars: 40 μm. (E) 10-week-old WT and Tg2 mice (n = 3 per group), as well as 3 month-old WT and dKO mice (n = 5 per group), were subjected to forced downhill running on a treadmill. Muscle performance was measured as time to exhaustion. Total running distance is also shown. *P < 0.05; ***P < 0.001.
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 6D). 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 6D). 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, 30). 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 6E), indicative of muscle weakness. dKO mice showed a more dramatic decrease in running capacity (Figure 6E). 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 mutation (9). 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 7, A and B). Furthermore, at least some of the intracellular dysferlin was colocalized with dynamin 2 in dKO muscle fibers (Figure 7A).
Intracellular accumulation of dysferlin in dKO and MCK-DNM2 transgenic mouse myofibers. (A) Immunostaining of TA muscle from WT and dKO mice to detect dynamin 2 and dysferlin. Intracellular accumulation of dysferlin was observed in dKO myofibers. Overlay images indicate localization of dynamin 2 and dysferlin in the intracellular aggregates in dKO muscle. Scale bars: 30 μm. (B) Immunostaining of TA muscle from WT and Tg2 mice to detect dysferlin. Intracellular accumulation of dysferlin was observed in Tg2 myofibers. Scale bars: 30 μm.
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 8, A and B). Soleus muscle of dKO mice showed a 2-fold increase in the number of type I fibers (Figure 8, A and B).
Control of skeletal muscle fiber type by miR-133a. (A) Metachromatic ATPase staining and anti–MHC-1 immunostaining of soleus muscle from WT and dKO mice at 12 weeks of age showed an increase in type I myofibers in dKO soleus muscle. H&E staining of the soleus muscles is also shown. Scale bars: 100 μm. (B) Percentage of type I myofibers in soleus muscles, determined by metachromatic ATPase staining. n = 6 (WT and dKO). (C) Expression of transcripts of MHC isoforms in soleus muscle, determined by real-time RT-PCR. n = 3 (WT and dKO). Expression of MHC isoforms from protein extracts of soleus, EDL, and TA muscles from WT and dKO mice was also determined by glycerol gel electrophoresis followed by silver staining.
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 8C). 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 8C). 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.