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. 2012 Mar 20;20(6):1222–1233. doi: 10.1038/mt.2012.35

Loss of miR-29 in Myoblasts Contributes to Dystrophic Muscle Pathogenesis

Lijun Wang 1,2, Liang Zhou 1,2, Peiyong Jiang 2,3, Leina Lu 2,3, Xiaona Chen 1,2, Huiyao Lan 2,4, Denis C Guttridge 1,5, Hao Sun 2,3,*, Huating Wang 1,2,*
PMCID: PMC3369280  PMID: 22434133

Abstract

microRNAs (miRNAs) are noncoding RNAs that regulate gene expression in post-transcriptional fashion, and emerging studies support their importance in a multitude of physiological and pathological processes. Here, we describe the regulation and function of miR-29 in Duchenne muscular dystrophy (DMD) and its potential use as therapeutic target. Our results demonstrate that miR-29 expression is downregulated in dystrophic muscles of mdx mice, a model of DMD. Restoration of its expression by intramuscular and intravenous injection improved dystrophy pathology by both promoting regeneration and inhibiting fibrogenesis. Mechanistic studies revealed that loss of miR-29 in muscle precursor cells (myoblasts) promotes their transdifferentiation into myofibroblasts through targeting extracellular molecules including collagens and microfibrillar-associated protein 5 (Mfap5). We further demonstrated that miR-29 is under negative regulation by transforming growth factor-β (TGF-β) signaling. Together, these results not only identify TGF-β–miR-29 as a novel regulatory axis during myoblasts conversion into myofibroblasts which constitutes a novel contributing route to muscle fibrogenesis of DMD but also implicate miR-29 replacement therapy as a promising treatment approach for DMD.

Introduction

microRNAs (miRNAs) are noncoding single-stranded RNAs of 21–25 nucleotides and constitute a novel class of gene regulators that are found in a variety of eukaryotic organisms. miRNAs negatively regulate their targets at the post-transcriptional level through binding to their 3′ untranslated regions (UTRs).1,2

With over 1,000 miRNA genes identified in the human genome, and a plethora of predicted messenger RNA (mRNA) targets, it is believed that these small RNAs add another layer of gene regulation that is subject to changes in human diseases. Altered expression of miRNAs has been discovered and identified to play key roles in the development of a variety of diseases, thus representing unique opportunities for therapeutic intervention.3,4,5 The therapeutic application of miRNAs involves two strategies. One strategy is directed toward a gain of function and aims to inhibit upregulated miRNAs by using miRNA antagonists, such as anti-miRs, locked nuclei acids (LNA), or antagomirs. The second strategy, miRNA replacement, involves the reintroduction of miRNA mimics to restore a loss of function.6

Muscular dysthophies are inherited disorders characterized by muscle degeneration and associated progressive wasting and weakness. Among these, Duchenne muscular dystrophy (DMD) is the most common lethal X-linked recessive disorder, affecting 1 in 3,500 live male births.7 Patients are usually confined to a wheelchair before the age of 12 and die in their late teens or early twenties. The origin of this disease stems from mutations in the dystrophin gene, which encodes a structural protein linking internal cytoskeleton to the extracellular matrix (ECM). The absence of dystrophin leads to sarcolemmar permeability, influx of calcium, and activation of proteases to cause myofiber necrosis and degeneration. This is followed to some extent by regeneration: muscle stems cells (satellite cells) are activated into muscle precursor cells (myoblasts) which are able to fuse together or with host myofibers to repair the damaged muscle. Unfortunately, complete regeneration is prevented by excessive synthesis and deposition of ECM proteins, which eventually lead to fibrosis.8,9,10 Thus, fibrosis is a prominent pathological hallmark of skeletal muscle in patients with DMD and contributes to progressive muscle dysfunction and the lethal phenotype of DMD.11,12

Fibrogenesis (the development of fibrosis) is a complex, incompletely understood process characterized by excessive accumulation of collagens and other ECM components. It is commonly held that fibrogenesis in dystrophic muscles is driven by the repeated bouts of muscle-fiber degradation and ensuing inflammation dominated by macrophages and T lymphocytes. The precise mechanisms leading to fibrosis in dystrophic muscle are not known but likely involve the action of several inflammatory cytokines, such as transforming growth factor-β (TGF-β).12 TGF-β is one of the most potent fibrogenic cytokines, and it contributes to the pathogenesis of a variety of fibrotic disorders including pulmonary fibrosis, cirrhosis, renal sclerosis, and scleroderma.13,14,15 Tissue fibrogenesis is mainly regulated by TGF-β1 isoform, which is upregulated in skeletal muscles of DMD patients and speculated to be a critical inducer of muscle fibrogenesis.16 However, the downstream signaling mediating the profibrogenic action of TGF-β is not fully explored.

At the cellular level, myofibroblasts are one of the key cellular components involved in tissue fibrosis,17 but little is known about the regulatory mechanisms of myofibroblast differentiation in muscle tissue. Although local resident fibroblasts are considered to be the main precursor cells, other cell types may also be the origin. For example, Li Y et al. demonstrated that after laceration-induced muscle injury, muscle progenitor cells are able to transdifferentiate into myofibroblasts and consequently contribute to the development of fibrosis,18 Interestingly, myoblasts cells from young and old mdx fibers showed elevated expression of collagen type I mRNA compared with the corresponding wild-type (WT) controls19; however, sufficient evidence is lacking to show that mdx myoblasts are indeed profibrogenic and their conversion into myofibroblasts may be a contributing route to muscle fibrogenesis in dystrophic muscles; furthermore, the molecular and cellular mechanisms underlying this fibrogenic conversion needs to be investigated.

Growing body of evidences suggests that miRNAs are involved in the process of fibrosis in several organs, including heart, lung, kidney, liver, and muscle (reviewed in ref. 20). In particular, a number of miRNAs, such as miR-2121,22 and miR-29 families,23,24 are emerging as common regulators of fibrogenesis in multiple tissues. Very recent findings implicated miR-29 family in cardiac, liver, pulmonary, skin, and muscle fibrosis through targeting ECM proteins such as collagens, fibrillins, and elastin.23,24,25,26,27 Most relevantly, Cacchiarelli D et al.27 demonstrated that intramuscular electroporation of an miR-29 expressing plasmid into 6-week-old mdx limb muscles could reduce the collagen and elastin mRNA expression, suggesting miR-29 as an antifibrotic molecule in dystrophic muscles. However, a striking feature of mdx skeletal muscle is that diaphragm and limb muscles have different fates. Limb muscles show a near-complete spontaneous resolution of inflammation with no significant fibrosis, whereas diaphragm displays persistent inflammation with progressive fibrosis, thus considered to be a more suitable target for antifibrotic research.12 Therefore, additional evidences are needed to support the antifibrotic function of miR-29. Furthermore, the effect of intramuscular delivery is limited to the local area and a more effective delivery approach is needed to achieve the systemic treatment effect.

Previously, our group identified miR-29 as a promyogenic factor during skeletal muscle cell differentiation.28,29 miR-29 promotes myogenesis by directly targeting Yin Yang 1 (YY1), a negative regulator of muscle genes. We further demonstrated that this regulatory circuit is disrupted in rhabdomyosarcoma which may contribute to the development of this tumor. These findings suggest that miR-29 involved circuitries are critical regulator of cellular differentiation and the disruption of this type of regulation may lead to the development of diseases. In this study, we investigated the role of miR-29 in DMD by using dystrophin-deficient mice. Our results elucidate the pleiotropic functions of miR-29 in both promoting muscle regeneration and inhibiting muscle fibrogenesis. Moreover, we show that local or systemic delivery of miR-29 mimics improved muscle regeneration and reduced fibrosis, implicating miR-29 replacement as a promising therapeutic approach in DMD.

Results

miR-29 is downregulated in mdx muscles

In order to gain insight into the function of miR-29 in DMD, we began by examining the expression levels of miR-29 in mdx mice, a widely used mouse model for DMD research. Total RNAs were extracted from multiple limb muscles including tibialis anterior (TA), gastrocnemius (Gas), and quadriceps (Quad) muscles from 4-week-old WT or mdx mice and subjected to quantitative reverse transcription-PCR (qRT-PCR) analysis for miR-29 expression. The results showed that all three members of miR-29 family, miR-29a, -29b, -29c, were downregulated in all three types of muscles (Figure 1a), with miR-29c displayed the most significant downregulation and will be the focus of this investigation henceforward. In addition, the downregulation of miR-29 was also detected in diaphragm and heart muscles (Figure 1b), further suggesting that miR-29 downregulation may play a critical role in mdx pathogenesis. In addition, miR-29 level seems to correlate strongly with disease severity, since considerably lower miR-29 expression was associated with muscles from utrophin-dystrophin double knock-out (DKO) mice, which, compared with mdx mice, more closely recapitulate the clinical signs and early lethality of DMD patients30; whereas heterozygous (Het) mdx mice displayed high miR-29 expression level (Figure 1c). Taken together, our data confirm that miR-29 is downregulated in dystrophic muscles, warranting the investigation of its function using mdx mice.

Figure 1.

Figure 1

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miR-29 is downregulated in mdx muscles. (a,b) The expressions of miR-29a, -29b, -29c in RNAs isolated from tibialis anterior (TA), gastrocnemius (Gas), quadriceps (Quad), diaphragm or heart muscles of wild-type (WT) or mdx mice. (c) The expressions of the above miRs in TA of WT, mdx, double knock-out (DKO) or heterozygous (Het) mice. Data are plotted as mean ± SD. *P < 0.05, **P < 0.01.

Restoration of miR-29 accelerates regeneration in mdx muscles

In response to the ongoing phase of degeneration in dystrophic muscles, cycles of regeneration attempt to compensate for the loss of muscle mass. Muscle regeneration is a complex process during which Pax7 positive quiescent satellite cells are activated into Pax7 and MyoD double positive muscle precursor cells (myoblasts) which are then able to fuse together or with host myofibers to regenerate the damaged muscle.31 The repaired and newly formed myofibers can be distinguished from undamaged fibers by their centrally located nuclei. Considering that myogenic differentiation represents an important step in the process of regeneration and miR-29 was known to act as a promoting factor during this process,28 we postulated that the downregulation of miR-29 might contribute to impaired regeneration capacity of dystrophic muscle. To test this notion, we employed gain-of-function approach by injecting 5 µmol/l of negative control (NC) or miR-29 mimics oligos into the TA muscles of 4-week-old mdx mice which are at the peak of massive degeneration-regeneration. Treatment was done three times a week for consecutive 2 weeks. Muscles were then collected 3 weeks after the initial injection. qRT-PCR analysis of miR-29 levels confirmed the successful restoration of miR-29 expression. A marked increase (~170 fold) of miR-29 was detected in miR-29 injected muscles as compared to NC oligos injected muscles (Supplementary Figure S1a). The degree of muscle regeneration was then assessed by multiple assays. Both hematoxylin and eosin staining and immunohistochemistry staining (IHC) for embryonic myosin heavy chain (E-MyHC) revealed an increased number of regenerating fibers in miR-29 injected muscles compared with NC muscles (Figure 2a). Scoring for E-MyHC–positive stained areas revealed a significant increase in miR-29 (10.2 ± 5%) injected muscles over that of muscles from NC (4 ± 2%, P < 0.001) (Figure 2b). The number of centrally located nuclei was also found to be significantly higher in miR-29 (285 ± 29) injected muscles than NC muscles (243 ± 41, P < 0.05) (Figure 2c). Furthermore, immunoblotting detected an increase of Pax7 (2.6-fold), MyoD (1.86-fold), and myogenin (1.3-fold) in miR-29 injected muscles (Figure 2d). Immunofluorescence (IF) staining for MyoD on the injected muscles confirmed that MyoD positive cell numbers increased by miR-29 injection (Supplementary Figure S1b). IF for Pax7 also revealed an increased number of Pax7+ cells in miR-29 treated muscles (Figure 2e,f). However, Pax7 was not predicted to be a direct target of miR-29 and reporter assay with Pax7-3′UTR fused downstream of luciferase (Luc) gene did not reveal an interplay between miR-29 and Pax7-3′UTR either (Supplementary Figure S1c,d). These findings indicate that miR-29 restoration enhanced myogenic differentiation. In addition, immunoblotting against YY1 revealed an obvious downregulation of YY1 (0.44-fold) (Figure 2d), suggesting that acceleration of regeneration by miR-29 could be mediated through downregulating YY1, a previously identified target of miR-29 during myogenic differentiation and the known repressor of myogenesis.28 Interestingly, we also observed a downregulation of macrophage marker, CD68, by miR-29 injection (Supplementary Figure S1e), suggesting miR-29 could also reduce macrophage infiltration, which may be an indirect effect of accelerated muscle repair. Taken together, these results support that miR-29 functions to promote regeneration in dystrophic muscles and implicate the potential use of intramuscular injection of miR-29 mimics as therapeutic approach for treating dystrophic muscles locally.

Figure 2.

Figure 2

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Loss of miR-29 in mdx myoblasts caused a defect on myogenic differentiation. (a) H&E and IHC staining for E-MyHC were performed on cryosections of mdx muscles injected with negative control (NC) or miR-29 mimics. n = 5 mice for each group. (b,c) Quantification of E-MyHC–positive fibers and fibers with centrally localized nuclei (CLN). (d) Left: immunoblotting for Pax7, MyoD, myogenin, and YY1 in the above muscles with GAPDH as a loading control. Results from three representative mice are shown. Right: quantification of western blots was done by densitometry. (e) IF staining for Pax7 was performed on the above NC or miR-29 injected muscles. (f) Positively stained cells were counted from at least 15 sections. E-MyHC, embryonic-MyHC; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; H&E, hematoxylin and eosin; IF, immunofluorescence; IHC, immunohistochemistry; YY1, Yin Yang 1. **P < 0.01, ***P < 0.001.

Myogenic differentiation is impaired in mdx myoblasts due to decreased miR-29 expression

Given that dystrophic muscle is composed of a mixture of multiple cell types including immune cells, muscle fibers, quiescent, and activated satellite cells, it is not clear at which cell compartment the downregulation of miR-29 is originated. Considering the known function of miR-29 in myoblast differentiation,28 we speculated that this downregulation occurs in myoblasts and leads to defective myogenic differentiation. To test this idea, primary myoblasts were freshly isolated from mdx or WT limb muscles and cultured as described before.32 Expectedly, miR-29 was found to be significantly downregulated in primary myoblasts from mdx muscles versus WT (Figure 3a). Furthermore, when induced to differentiation, mdx myoblasts displayed a delay in myogenic program as assessed by a reduction of both MyHC positive myotube formation (Figure 3b) and decreased expressions of myofibrillar genes, MyHC IIB and troponin over a period of 48 hours (Figure 3c). To test whether this defect is due to the loss of miR-29, we found that restoration of miR-29 levels by oligo transfection (Figure 3d, >500-folds) augmented the myogenic differentiation as evidenced by an increase of myotube formation (Figure 3e) as well as the RNA expression levels of myofibrillar genes (Figure 3f). Furthermore, transfections with MyHC or troponin luciferase reporters (MyHC-Luc and TnI-Luc) in mdx primary myoblasts showed that myogenic activity was substantially elevated with miR-29 restoration (Figure 3g, P < 0.05). Collectively, the above results suggest that loss of miR-29 in mdx primary myoblasts leads to a defect on myogenic differentiation which likely contributes to the impaired regeneration capacity in dystrophic muscles.

Figure 3.

Figure 3

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miR-29 is downregulated in mdx myoblasts. (a) Expression of miR-29 in primary myoblasts from WT or mdx muscles. (b) Left: primary myoblasts from mdx muscles were kept growing (DM 0 hour) or differentiated (DM) for 7, 24, or 48 hours at which times cells were immunostained for MyHC. Right: positively stained cells were quantified. Numbers indicate the average number of MyHC positive cells counted from a minimum of 10 randomly chosen fields. Graphs are plotted as mean ± SD. Images were taken at 24 hours. (c) Expressions of MyHC or troponin RNAs in WT or mdx myoblasts differentiated for the indicated times. (d) Expression of miR-29 in mdx primary myoblasts transfected with NC or miR-29 oligos. (e) Left: the above transfected cells were differentiated for 48 hours at which time the cells were photographed under phase contrasts or immunostained for MyHC. Right: positively stained cells were quantified as in b. (f) Expressions of MyHC and troponin RNAs in NC or miR-29 transfected cells at different time points of differentiation. (g) mdx myoblasts were transfected with MyHC-Luc or TnI-Luc reporter plasmids and NC or miR-29 oligos. Cells were then differentiated for 48 hours at which time luciferase activities were determined. The data represent the average of three independent experiments ± SD. *P < 0.05, **P < 0.01. DM, differentiation medium; NC, negative control; WT, wild-type.

Intramuscular injection of miR-29 inhibits ECM gene expression in mdx muscles

In addition to its role in muscle regeneration as suggested by the above findings, we were intrigued to explore the additional events that miR-29 could be involved in. A genome-wide transcriptome analysis was thus conducted to globally characterize miR-29–mediated transcriptome changes using a high throughput mRNA sequencing (mRNA-seq) platform which provides higher level of accuracy and broader dynamic ranges compared to traditional microarray-based platform.33 Total RNAs were extracted from mdx limb muscle injected with miR-29 or NC oligos as above and subjected to mRNA-seq using Illumina GAIIx platform. A total of 15 million and 17 million raw reads were sequenced from miR-29 and NC samples, respectively, which were then mapped to reference genome NCBIM37.61 via Tophat v1.2.34 The majority of reads can be mapped to exonic regions (>5 fragments per kilobase of transcript per million fragments mapped (FPKM)) and much fewer (~0.2 FPKM) in introns and intergenic regions (Supplementary Figure S2), indicating great specificity for expressed mRNA and rejection of genomic DNA and unspliced pre-mRNA. Cufflinks.cuffdiff program (v1.0.0) 35 was subsequently employed to identify the differentially expressed genes under a false discovery rate of 5%. As a result, a total of 476 and 1,952 genes were found to be up- and downregulated in miR-29 injected muscles versus NC muscles (Figure 4a and Supplementary Tables S1 and S2). Subsequent Gene Ontology (GO) analysis with upregulated list of genes revealed that the top ranked lists of enriched GO categories include “sarcomere”, “contractile fiber part”, “myofibril”, “contractile fiber”, “I band”, “Z disc” (Supplementary Table S3a), which is in agreement with the afore-identified roles of miR-29 in accelerating muscle regeneration. Strikingly, GO analysis with downregulated list of genes revealed an over-representation of ECM genes presented in GO categories such as “extracellular matrix”, “extracellular matrix part”, “extracellular region part”, “collagen” et al. (Figure 4b and Supplementary Table S3b). This is in agreement with the emerging reports demonstrating the pivotal role of miR-29 in ECM remodeling as well as fibrosis of multiple tissues.23,24,25,26

Figure 4.

Figure 4

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Overexpression of miR-29 in mdx muscles downregulates fibrotic genes. (a) Differentially expressed genes in mdx muscles injected with NC and miR-29 oligos as determined by mRNA-seq. X- and Y-axis represent the log2 based FPKM values for expressed genes in NC and miR-29 samples, respectively. (b) Over-represented GO terms by GO analysis of downregulated list of genes. BP, biological process; CC, cellular component; KEGG, Kyoto Encyclopedia of Genes and Genomes; SP_PIR, a database of protein super-family names. (c) Coverage plot showing a 56-kb region encompassing the ACTA2 (α-SMA) gene on chromosome (Chr) 19; the gene structure is shown in blue below the graph. (d) Expressions of collagen 1A1 (Col 1A1), collagen 1A2 (Col 1A2), collagen 3A1 (Col 3A1), α-smooth muscle actin (α-SMA) or vimentin (VIM) in NC or miR-29 injected muscles. (e) Expressions of the above genes in TA muscles injected with negative control (Anti-NC) or miR-29 inhibitor oligos (Anti-miR-29). FPKM, fragments per kilobase of transcript per million fragments mapped; GO, gene ontology; mRNA, messenger RNA; NC, negative control; TA, tibialis anterior.

In support of the GO analysis, the expressions of several fibrotic markers: collagen 1A1 (Col 1A1), collagen 1A2 (Col 1A2), and collagen 3A1 (Col 3A1), α-smooth muscle actin (ACTA2 or α-SMA) and vimentin (VIM) were found to be downregulated in miR-29 injected muscles by both mRNA-seq (Figure 4c, α-SMA expression) and qRT-PCR analysis (Figure 4d). To substantiate the above findings, loss-of-function experiment was performed by injecting anti-miR-29 LNA oligos into mdx TA muscles. As opposite to the effect of miR-29 mimics, anti-miR-29 injection increased the expression levels of all the above fibrotic markers as compared to negative control (anti-NC) oligos (Figure 4e). Together, these data suggest that the decrease of miR-29 in mdx muscles may contribute to muscle fibrogenesis and warrant further investigation of the underlying molecular and cellular mechanisms.

Loss of miR-29 promotes myoblasts conversion into myofibroblasts

Knowing that loss of miR-29 occurs in myoblast compartment, we speculated that it induces elevation of collagen expression and transdifferentiation of myoblasts into myofibroblast, which contributes to muscle fibrogenesis in dystrophic muscles. In agreement with this thinking, myoblasts isolated from mdx muscle showed increased fibrogenic potential as evidenced by increased expressions of Col 1A1, Col 1A2, and Col 3A1 mRNAs (Figure 5a) whereas their myogenic differentiation is suppressed (Figure 3). As expected, miR-29 restoration caused an inhibition of collagen expression (Figure 5b), suggesting that miR-29 targeted collagens. This notion was further examined by using reporters with a fragment of the Col 1A1, Col 1A2, or Col 3A 3′ UTR containing the miR-29 binding site fused downstream of the firefly Luc gene. Cotransfections of the reporter plasmid (WT) with miR-29 caused significant repressions of Luc activities (Figure 5c). In addition to collagen, a dozen of other ECM genes were downregulated by miR-29 injection (Supplementary Table S2). Among them, Mfap2 and Mfap5 (microfibrillar-associated protein-2 and -5) are ECM proteins, which are localized with microfibrils in elastin network.36 Interestingly, a miR-29 target site was found on the 3′UTR region of Mfap5 (Figure 5d). To test whether Mfap5 represents a novel target of miR-29, the predicted binding sites of miR-29 was cloned into a pMIR-reporter vector. Cotransfection of the resultant reporter plasmids together with miR-29 into C2C12 cells caused a repression on Luc activities of the reporter (Figure 5e), indicating that miR-29 indeed directly targets Mfap5. Analogous to collagen expressions, myoblasts isolated from mdx muscle showed increased Mfap5 expression (Figure 5f) and restoration of miR-29 decreases Mfap5 expression (Figure 5g). Consistently, intramuscular injection of anti-miR-29 into mdx muscle increased its expression (Figure 5h). Collectively, our findings suggest that high level of miR-29 is important for driving myogenic differentiation and loss of miR-29 promotes transdifferentiation of myoblasts into myofibroblasts, which may be attributable to ECM accumulation and muscle fibrogenesis in dystrophic muscles. Lastly, to prove that the transdifferentiation indeed occurs in vivo, IF staining uncovered a large number of α-SMA+ cells in mdx muscles (Figure 5i, arrows) which was not seen in WT muscles undergoing cardiotoxin-induced regeneration (data not shown). Moreover, some of the α-SMA positive cells co-stained with MyoD (Figure 5i, asteroid), indicating that these cells are likely to be of myoblast origin. Consistently, myoblasts isolated from mdx co-stained for α-SMA and MyoD in the culture (Supplementary Figure S3). Remarkably, miR-29 restoration in vivo significantly reduced the percentage of MyoD+/α-SMA+ cells (Figure 5j), consistent with the idea that miR-29 inhibits myoblast transdifferentation. These in vivo data add strong evidence to support our thinking that myoblasts conversion into myofibroblasts represents a previously unidentified contributing route to muscle fibrogenesis.

Figure 5.

Figure 5

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Loss of miR-29 promotes myoblasts transdifferentiation into myofibroblasts. (a) Expressions of Col 1A1, Col 1A2, and Col 3A1 in primary myoblasts freshly isolated from WT or mdx muscles. (b) Primary myoblasts from mdx muscles were transfected with NC or miR-29 oligos and examined for Col 1A1, Col 1A2, and Col 3A1 expressions. (c) WT or mutant Col 1A1, Col 1A2, or Col 3A1-3′UTR luciferase reporter constructs were transfected into mdx primary myoblasts with NC or miR-29 oligos. Luciferase activities were determined at 48 hours post-transfection. Relative luciferase unit (RLU) is shown with respect to NC cells where normalized luciferase values were set to 1. The data represents the average of three independent experiments ± SD. (d) Predicted binding between mmu-miR-29c and mouse 3′UTR of Mfap5. (e) WT or mutant Mfap5-3′UTR luciferase reporter constructs were transfected into C2C12 cells with NC or miR-29 oligos. Luciferase activities were determined as above. (f) Expressions of Mfap5 in primary myoblasts from WT or mdx muscles. (g) Primary myoblasts from mdx muscles were transfected with NC or miR-29 oligos and examined for Mfap5 expression. (h) Expressions of the Mfap5 mRNAs in mdx TA muscles injected with negative control (Anti-NC) or miR-29 inhibitor oligos (Anti-miR-29). (i) IF staining for MyoD (green) and α-SMA (red) were performed on cryosections of mdx TA muscles. DAPI (blue) staining was also performed to visualize the nuclei. A field with three types of cells is shown on the left: arrows, MyoD-/α-SMA+ cells; arrowhead, MyoD+/α-SMA- cells; asterisk, MyoD+/α-SMA+ cells. Higher magnification of one of the MyoD+/α-SMA+ cells is presented on the right. (j) TA muscles from mdx mice were injected with NC or miR-29 mimics oligos. Quantification of the MyoD+/SMA+ cells on the above muscles were performed on a minimal of 15 sections for each group. N = 5 mice per group. Data are plotted as mean ± SD.*P < 0.05. α-SMA, α-smooth muscle actin; DAPI, 4′,6-diamidino-2-phenylindole; mRNA, messenger RNA; NC, negative control; TA, tibialis anterior; UTR, untranslated region; WT, wild-type.

miR-29 is downregulated by TGF-β signaling

Having gained insights into the role of miR-29 during the conversion of myoblasts to myofibroblasts, we now turned our attention to its upstream regulator by asking: what leads to the downregulation of miR-29 in fibrotic muscles? It is known that TGF-β is highly expressed in dystrophic muscles,16,37,38 likely playing a key role in the initiation of fibrotic cascade. In addition, several recent reports demonstrated the regulation of miR-29 by TGF-β in various tissue fibrosis.39,40,41 We thus speculated that downregulation of miR-29 in dystrophic muscles could be caused by elevated TGF-β level, and the profibrogenic action of TGF-β mediated through miR-29 could represent a novel signaling event contributing to muscle fibrogenesis.

Consistent with previous report showing TGF-β was elevated in dystrophic muscles,16,37,38 qRT-PCR analysis detected increased TGF-β mRNA expressions at all ages of mdx muscles (Figure 6a). This is correlated with the downregulation of miR-29 (Figure 1 and Supplementary Figure S4). We then sought to downregulate TGF-β signaling by intramuscular injection of decorin which was proven to inhibit TGF-β activation and thought to be an effective antifibrotic agent.38 To our expectation one dose injection of 10 mg/ml decorin resulted in significant upregulation of miR-29, whereas collagen and Mfap5 mRNAs were downregulated (Figure 6b,c). This suggests that elevated TGF-β signaling caused the suppression of miR-29 and consequent upregulation of ECMs. Furthermore, TGF-β treatment of mdx myoblasts led to strong inhibition of miR-29 expression but an increase on collagen and Mfap5 expressions (Figure 6d,e). Together, these findings suggest a functional existence of TGF-β regulation on miR-29 in promoting fibrogenic differentiation of mdx myoblasts.

Figure 6.

Figure 6

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Elevated TGF-β signaling downregulates miR-29 in dystrophic muscles and mdx myoblasts. (a) Expressions of TGF-β mRNAs in TA muscles of WT or mdx mice at indicated ages. (b) Expressions of miR-29 in TA muscles of mdx mice injected with decorin (+) or PBS control (-). (c) Expressions of collagens and Mfap5 in TA muscles of mdx mice injected with decorin (+) or PBS control (-). (d,e) Primary myoblasts from mdx muscles were treated with or without TGF-β and examined for expressions of miR-29 or collagens and Mfap5. mRNA, messenger RNA; PBS, phosphate-buffered saline; TA, tibialis anterior; TGF-β, transforming growth factor-β WT, wild-type.

Systemic delivery of miR-29 into mdx mice reduces fibrosis

Having gained a picture of how miR-29 functions to promote regeneration and to inhibit fibrogenesis as well as its underlying molecular and cellular mechanisms, we attempted to exploit miR-29 mimics as an antifibrotic agent in dystrophic muscles. Although the results in Figure 3 demonstrated that the intramuscular injection of miR-29 mimics into limb muscles caused dramatic impact on the transcriptome and downregulated fibrotic markers, the effect is nevertheless limited to injected areas only; the most severely affected diaphragm or heart muscles will not be reachable by this approach. Trichrome staining on cryofixed muscle sections revealed that collagen deposition can be observed in mdx diaphragm starting 6-week-old and worsened dramatically in older mice (Supplementary Figure S4a). Conversely, no detectable collagen deposition was observed in WT mice of the same age.

In order to achieve delivery into diaphragm muscles, a systemic delivery through tail vein injection was tested on 9-month-old mdx mice. Naked miRNA oligos are rapidly degraded in biofluids, and thus, we formulated the miRNAs in a neutral lipid emulsion delivery vehicle. The formulations were administered intravenously by tail vein injections. Each dose contained 44 µg of formulated oligos, which equals 14–15 mg/kg per mouse with an average weight of 30 g. Three doses were given following the scheme in Figure 7a and the diaphragm muscles were collected for staining at the end of 21 days. As controls, a separated group of animals were injected with vehicle formulated with NC oligos. Successful delivery into diaphragm muscles was confirmed by detecting fluorescein isothiocyanate (FITC) dye using a live imaging system (Supplementary Figure S4b and Supplementary Materials and Methods). Restoration of miR-29 levels was found in multiple tissues with a marked increase of ~50-folds in diaphragm muscles by qRT-PCR (Figure 7b). In-situ hybridization with a digoxigenin (DIG)-labeled miRNA LNA probe also detected miR-29 signals in various tissues, demonstrating successful delivery of oligos with neutral lipid emulsion (Supplementary Figure S5c and Supplementary Materials and Methods). To assess the degree of fibrosis, trichrome staining was performed on frozen sections of diaphragm muscles and the results revealed a strong decrease (50%, P < 0.01) of positively stained areas in the miR-29 injected group (Figure 7c). The results from IHC and IF staining for collagen 1 proteins also confirmed a reduction of collagen 1 expressions in the miR-29 injected group (Figure 7d, 37% reduction, P < 0.01 and Figure 7e). In addition to an evident reduction of fibrosis, regeneration was found to be improved in miR-29 treated group as revealed by an elevated expression of Pax7, MyoD, and myogenin as well as downregulation of YY1 (Figure 7f). As a result, the overall morphology was found to be markedly improved as less damaged area (a sum of fibrotic and fatty depositions, 13% reduction, P < 0.0001) was observed in miR-29 treated group as revealed by hematoxylin and eosin staining (Figure 7h). Taken together, these findings implicate that systemic administration of miR-29 mimics decreases the existing fibrosis and improves muscle repair, pointing to miR-29 mimics as a potential treatment approach for DMD.

Figure 7.

Figure 7

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Systemic delivery of miR-29 oligos reduces fibrosis in mdx diaphragm. (a) Administration scheme for miR-29 injection. NC or miR-29 oligos formulated with liposome were injected into mdx mice through tail vein at day (d) 0, 4, and 7. Mice were killed and diaphragm muscles were harvested at day 21 for histology and immunostaining. n = 5 mice for each treatment group. (b) Expressions of miR-29 in various tissues collected at day 3 after the injection. (c) Trichrome staining of the fibrotic areas in NC and miR-29 injected diaphragm muscles. The positively stained areas were quantified with Image-Pro Plus from a minimum of 15 sections. (d) IHC staining of the above muscles with collagen 1. The positively stained areas were quantified as the above. (e) IF staining of the above muscle with collagen 1. (f) Expressions of Pax7, MyoD, myogenin, and YY1 mRNAs in the above treated diaphragm muscles. (g) H&E staining of the above NC or miR-29 injected mdx muscles. (h) Damaged areas were identified as fibrotic and fatty areas and quantified as the above. **P < 0.01, ***P < 0.001. H&E, hematoxylin and eosin; IF, immunofluorescence; IHC, immunohistochemistry; mRNA, messenger RNA; NC, negative control; SI, small intestine; TA, tibialis anterior; YY1, Yin Yang 1.

Discussion

In the current study, we present evidences for the pleiotropic roles of miR-29 in DMD physiopathology. Downregulation of miR-29 in dystrophic muscles not only results in impaired regeneration but also contributes to muscle fibrogenesis. Although miR-29 was implicated in regulating fibrosis in mdx muscles by Cacchiarelli D et al.,27 our study for the first time uncovers a novel cellular mechanism. We demonstrate that the downregulation of miR-29 stems from myoblast compartments, which causes suppression of myogenic differentiation and promotion of fibrogenic transdifferentiation. In lines with our previous findings,28 YY1 is likely the downstream mediator of miR-29 effect on myogenic regeneration. In addition, we added one more target, Mfap5, to the growing list of ECM molecules regulated by miR-29. We showed that both Mfap5 and multiple collagen genes were direct targets of miR-29 in mediating its role in muscle fibrogenesis. This is consistent with emerging results implicating miR-29 as a critical mediator in organ fibrosis including lung, hepatic as well as kidney fibrosis.20 However, we cannot exclude the possibility that downregulation of miR-29 also occurs in other cell compartments such as quiescent satellite cells, infiltrating immune cells, fibroblasts, as well as myofibers. As revealed by genome-wide transcriptome profiling, miR-29 injection caused a drastic change on transcriptome in mdx muscles. Other than ECM genes, GO analysis with downregulated genes also revealed an enrichment for GO terms including “antigen processing and presentation”, “immune effector process”, “positive regulation of immune response”, “positive regulation of immune system process”, “immune response” et al. (Figure 4c and Supplementary Table S3b), strongly suggesting that miR-29 may also be involved in modulating inflammatory events. This is further supported by recent findings showing the expression of miR-29 in immune cells and its function in modulating immune responses42,43

Resident cells, infiltrating inflammatory cells, and ECM components within dystrophic muscle tissue create a complex signaling environment. Our findings for the first time provide a full spectrum of miR-29 influence on dystrophic muscles and implicate miR-29 as a multifaceted molecule involved in diverse molecular processes and signaling pathways. Further elucidation of its modes of action in various cellular compartments will unveil novel mechanisms of muscle dystrophinopathy.

Fibrosis is a prominent pathological feature of skeletal muscle in patients with DMD. It is believed that fibroblasts are the major source of ECM production. However, the contribution from other sources has long been speculated. The transdifferentiation of myoblasts into myofibroblasts has been nicely demonstrated in C2C12 culture, which undergoes ECM remolding and fibrogenic conversion upon TGF-β treatment. The underlying mechanisms mediating the profibrogenic effect of TGF-β in C2C12 cells were not fully understood. In one case, TGF-β was able to induce Smad-dependent upregulation of spiningosin kinase SK1 in C2C12 myoblasts. Rho kinase signaling also appeared to be implicated in the TGF-β–mediated transition of myoblasts into myofibroblasts downstream of SK1 activation.40 Similarly, downregulation of Notch2 expression has also been linked to nonmuscle fibrotic tissue and TGF-β–dependent induction of myofibroblast makers in C2C12 myoblasts.44 Here in the current investigation, we present evidences to show that mdx myoblasts indeed possess high propensity of transdifferentiation into myofibroblasts whereas its myogenic potential is defective compared to WT. Loss of miR-29 through elevated TGF-β signaling likely contributes to this intrinsic defect of mdx myoblast. Thus, TGF-β–miR-29 axis represents a novel circuit that is critical in determining the fate of myoblasts: the high level of TGF-β in dystrophic muscles causes a loss of miR-29 and subsequent conversion of myoblasts into myofibroblasts as well as overproduction of collagens, which contributes to muscle fibrogenesis. Therefore, our findings reinforced the potent roles of TGF-β in inducing muscle fibrogenesis and uncover a novel mechanism mediating its action. Despite the mounting evidences supporting the antifibrotic roles of miR-29, the underlying mechanism was not fully explored. Our findings thus identify a novel cellular mechanism underlying miR-29 action in tissue fibrosis.

The rational for developing miRNA therapeutics is based on the premise that aberrantly expressed miRNAs play key role in the development of human diseases, and that correcting these deficiencies by either antagonizing or restoring miRNA function may provide a therapeutic benefit. Although miRNA antagonists are conceptually similar to other inhibitory therapies, restoring a function of miRNA by replacement is less characterized approach. Ectopic expression of one single miRNA to impact a disease was nicely demonstrated in human cancers. Systemic delivery of synthetic miR-16 inhibited the growth of metastasis prostate tumors45; similarly, systemic delivery of miR-34 blocks tumor growth in mouse models of non-small cell lung cancer.46 In both studies, administration of synthetic miRNA mimics formulated with lipid-based delivery vehicles were found to be well-tolerated by mice thus representing a highly effective and nontoxic treatment approach for human cancers. Here we present the first line of evidence to demonstrate that miR-29 mimics are effective in improving pathology of dystrophic muscle. Not only it led to remarkable reduction of existing fibrosis but also improved regeneration and decreased damaged areas.

Most of the interventions to test antifibrotic therapies in mdx mice started before the morphological onset of diaphragm fibrosis. Our findings, however, identified a therapy that can reverse the existing muscle fibrosis. In our study, no obvious adverse effect was observed after treatment with miR-29 mimics although more extensive and long-term toxicity studies are required. Intramuscular administration of miR-29 mimics led to 50–200-fold restoration of miR-29 level, thus representing a promising strategy for treating injured muscles locally. However, systemic delivery is necessary to reach less accessible tissues such as diaphragm and heart. Compared to local injection, the overall abundance of systemically delivered miR-29 oligos in diaphragm and limb muscles was less (Figure 7b). Nevertheless, significant effect was achieved, suggesting that only minimal amounts of miR-29 are needed to elicit a therapeutic effect.

Taken together, our results identified miR-29 as a pleiotropic molecule in DMD. As modeled in Figure 8, we propose that downregulation of miR-29 in dystrophic muscles contributes to dystrophic pathogenesis by inhibiting regeneration and stimulating fibrogenesis. Detailed mechanistic studies uncovered a novel mechanism underlying miR-29 action in dystrophic muscles. Our results elucidated that miR-29 is downregulated in myoblasts which leads to inhibition of myogenic differentiation and increased propensity of transdifferentiation into myofibroblasts. Further investigation showed that miR-29 expression is inhibited by TGF-β signaling during fibrogenic conversion. Finally, we demonstrated that systemic delivery of miR-29 mimics led to significant improvement of diaphragm muscle pathology by reducing existing fibrosis and increasing regeneration, suggesting that miR-29 replacement therapy can be a novel treatment approach for DMD.

Figure 8.

Figure 8

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A model of the role of miR-29 in dystrophic muscles. The model depicts the critical roles of miR-29 in muscle regeneration and muscle fibrogenesis. The model depicts the roles of the TGF-β–miR-29 regulatory axis in transdifferentiation of myoblasts into myofibroblasts. In the normal myogenesis, NF-κB-YY1 regulated miR-29 promotes the differentiation through downregulating YY1, which leads to successful myogenic differentiation and muscle regeneration. In dystrophic muscles, miR-29 is downregulated by increased TGF-β signaling, leading to overexpression of ECM genes including collagens, which drives the conversion of myoblasts into myofibroblasts thus contributing to muscle fibrogenesis and defective regeneration. Arrows: activation; blunted arrows: repression. ECM, extracellular matrix; MFAP, microfibrillar-associated protein; NF-κB, nuclear factor-κB; TGF-β, transforming growth factor-β YY1, Yin Yang 1.

Materials and Methods

Animal studies. Mdx (C57BL/10 ScSn DMDmdx) were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice were housed in the animal facilities of The Chinese University of Hong Kong (CUHK) under conventional conditions with constant temperature and humidity and fed a standard diet. Animal experimentation was approved by the CUHK Animal Ethics Committee. For intramuscular injection of miRNA oligos, ~3- to 4-week old mdx mice were injected with miR-29c mimics (Life Technologies, Carlsbad, CA) or scramble control oligos (Ambion) into the left and right TA muscle, respectively. The oligos were prepared by incubating 6 µl of 50 µmol/l pre-miR oligos with 6 µl of Lipofectamine 2000 for 15 minutes and a final volume of 60 µl in OPTI-EM (Life Technologies) were injected into the muscle. Injections were administrated every other day for 2 weeks. Muscles were harvested at the end of third week, and frozen sections prepared for histology analysis. Proteins and RNAs were extracted for western blot analysis and real-time RT-PCR analysis. Five mice were used in each group. For decorin studies, ~6-week-old mdx mice were injected with 30 µl of decorin at 10 mg/ml into the TA muscle. Muscles were harvested 7 days later, and RNAs were extracted for qRT-PCR analysis. For systemic delivery of miR-29 mimics, 9-month-old mdx mice were injected with miR-29c mimics or NC oligos (RiboBio, Guangzhou, China). Oligos were prepared by formulating with Max Suppressor In Vivo RNA-LANCER-II according to manufactory (Bioo Scientific, Austin, TX). Injections were performed for three times at day 0, 4, and 7. Diaphragm muscles were collected at day 21 for histology analysis as well as protein and RNA measurements. For miR-29 expression analysis, various tissues were collected at day 3 after the initial injection and used for qRT-PCR analysis.

Cell. Mouse C2C12 myoblasts were obtained from American Type Culture Collection (Manassas, VA) and cultured in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum, 2 mmol/l L-glutamine, 100 U/ml penicillin, and 100 µg of Streptomycin at 37 °C in 5% CO2. For myogenic differentiation, cells were seeded in 60 mm or 100 mm plates and when 90% confluent they were shifted to Dulbecco's modified Eagle's medium without fetal bovine serum containing 2% horse serum. Cells were incubated with various doses of TGF-β1 (R&D systems, Minneapolis, MN).

Primary myoblast isolation and culture. Myoblasts were isolated from ~1-week-old C57/BL10 or mdx muscles adopted from the described procedures (Rando and Blau, 1994). Briefly, total hind limb muscles (3–6 mice per group) were digested with type IV collagenase (Life Technologies; 5 mg/ ml) and dispase II (Life Technologies; 1.4 mg/ml) for 0.5 hour, and cell suspensions were further homogenized by pipetting before being filtered through 70 and 40 µm filters. The obtained cells were pre-plated on uncoated cell culture plates in F10 media (Life Technologies) to selectively enrich for myoblasts. After two rounds of pre-plating, the cell suspension was plated on Gelatin-coated plates (Iwaki, Tokyo, Japan) in F10 medium (Life Technologies) supplemented with 20% fetal bovine serum and Basic Fibroblast Growth Factor (Life Technologies; 25 ng/ml). Primary myoblasts were used at passage 3–5 after isolation.

Transfections and infections. Transient transfections with miRNA precursor oligos and siRNA oligos or DNA plasmids were performed in 60 mm or 100 mm dishes with Lipofectamine reagent as suggested by the manufacturer (Life Technologies). For Luc experiments, C2C12 and primary myoblasts were transfected in 12-well plates. Cell extracts were prepared and Luc activity was monitored as previously described47 or using dual-luciferase kit (Promega, Madison, WI).

Oligonucleotides. Precursor miRNA oligos were obtained from Ambion. Mercury LNA microRNA or control oligos were obtained from Exiqon (Woburn, MA). In each case 50 µmol/l oligos were used for transient transfections into cells. For systemic delivery, the miR-29c oligos with the modifications of methylation, thio-, cholesterol were obtained from RiboBio.

DNA constructs. Col 1A1-3′-UTR, Col 1A2-3′-UTR, Col 3A1-3′-UTR Luc reporter plasmids were kind gifts from Dr Ahlquist Paul.48 For the construction of mutant plasmid, the 29 base pair seed region of the predicted miR-29 binding site was deleted from the above parental constructs using QuickChange XL-mutagenesis (Stratagene, La Jolla, CA). MyHC-Luc and Troponin-Luc were used as described.49 Renilla Luc reporter was obtained from Promega and used according to manufactory. To construct Mfap5-3′UTR reporter plasmids, a 45 bp fragment encompassing miR-29 binding site was cloned into pMIR-report vector (Life Technologies) between Spa1 and Sac1 sites. Mutant reporter plasmids were generated by mutating the seed region from TGGTGCT to TACCTCT. A Pax-3′UTR Luc reporter was generated and used as described.50

RT-PCR and real-time RT-PCR. Total RNAs from cells were extracted using TRIzol reagent (Invitrogen). Expression of mature miRNAs was determined using the miRNA-specific Taqman microRNA assay kit (Life Technologies) using ABI PRISM 7900HT Sequence Detection System (Life Technologies). U6 was used for normalization. Expression of mRNA analysis was performed with SYBR Green Master Mix (Applied Biosystems) as described using GAPDH for normalization.49

Immunoblotting, immunostaining, and IHC. For western blot analysis, total cell extracts were prepared and used as previously described.47 The following dilutions were used for each antibody: Pax 7 (Developmental Studies Hybridoma Bank; 1:2,000), MyoD (Santa Cruz Biotechnology, Santa Cruz, CA; 1:2,000), myogenin (Santa Cruz Biotechnology; 1:2,000), YY1 (Santa Cruz Biotechnology; 1:2,000), troponin (Sigma-Aldrich, St Louis, MO; 1:2,000), MyHC (Sigma-Aldrich; 1:2,000), α-tubulin (Sigma-Aldrich; 1:5000), and GAPDH (Santa Cruz Biotechnology; 1:5,000). IF of cultured cells was performed using the following antibodies: MyHC (Sigma-Aldrich; 1:350), α-SMA (Millipore, Billerica, MA; 1:400), MyoD (DAKO, Carpinteria, CA; 1:400). Frozen muscle sections were prepared and stained as previously described. IF on frozen muscle sections was performed using the following antibodies: collagen (Novus Biologicals, Littleton, CO; 1:200), α-SMA (Sigma-Aldrich; 1:200), MyoD (Santa Cruz Biotechnology; 1:100); IHC on frozen muscle sections was performed using an antibody against E-MyHC (Novocastra; Leica Microsystems, Wetzlar, Germany) at a dilution of 1:200; For E-MyHC and CNL quantification, counts were performed from a minimum of 20 randomly chosen fields, from 5–6 sections throughout the length of the muscle in 4–6 per group. Hematoxylin and eosin staining on frozen muscle sections were performed as previously described.35 Masson's trichrome staining was performed according to the manufactory (ScyTek Laboratories, Logan, UT). The quantification of trichrome or IHC stained areas and damaged areas is processed with Image-Pro plus software (Media Cybernetics, Bethesda, MD) from a minimum of 15 images. All fluorescent images were captured with a confocal laser scanning microscope (FV1000; Olympus, Tokyo, Japan). All samples were imaged with the 20× or 40× objective lens. Pictures were captured in Kahlman frame giving an average of two scans using the Olympus microscope FV1000 and the accompanying software FV10-ASW (version 01.07.02.02; Olympus).

Sequencing and base calling. Preparation of transcription libraries for sequencing on the Illumina GA2x platform was carried out using the mRNA-seq sample preparation kit (Illumina, San Diego, CA) (Part no. 1004898 Rev. D) according to the manufacturer's standard protocol. Briefly, total RNA (10 µg) was subjected to two rounds of hybridization to Oligo (dT) beads. The resulting mRNAs were fragmented via incubation for 5 minutes at 94 °C with the Illumina-supplied fragmentation buffer. The first strand of complementary DNA was next synthesized by reverse transcription using random primers. Second-strand synthesis was conducted by incubation with RNase H and DNA polymerase I. The resulting double-stranded DNA fragments were subsequently end-repaired, and A-nucleotide overhangs were added by incubation with Taq Klenow lacking exonuclease activity. After the attachment of anchor sequences, fragments were PCR-amplified using Illumina-supplied primers and loaded onto the GA2x flow cell. DNA clusters were generated with an Illumina cluster station with Paired-End Cluster Generation Kit v2 (Illumina), followed by 51 × 2 cycles of sequencing on the GA2x (Illumina) with Sequencing Kit v3 (Illumina). Genome Analyzer Sequencing Control Software (SCS) v2.5, which could perform real-time image analysis and base calling, was used to carry out the image processing and base calling during the chemistry and imaging cycles of a sequencing run. The default parameters within the data analysis software (SCS v2.5) from Illumina were used to filter poor-quality reads. In the default setting, a read would be removed if a chastity of less than 0.6 is observed on two or more bases among the first 25 bases.

Read mapping to genome with splice-aware aligner. Sequenced fragments were mapped to reference genome NCBIM37.61 using TopHat version 1.2. Cufflinks version 1.0.0 was then used to estimate transcript abundances of RNA-Seq experiments. Abundances were reported in FPKM which is conceptually analogous to the reads per kilobase per million reads mapped (RPKM) used for single end RNA-seq.

Statistical analysis. Statistical significance was assessed by the Student's t-test. (*P < 0.5; **P < 0.01; ***P < 0.001)

SUPPLEMENTARY MATERIAL Figure S1. Intramuscular injection of miR-29 oligos restores miR-29 expression in mdx limb muscles and improved regeneration. Figure S2. RNA-seq reveals that miR-29 downregulates fibrotic genes in mdx muscles. Figure S3. Co-staining of MyoD and α-SMA in mdx myoblasts. Figure S4. miR-29 is downregulated in dystrophic muscles. Figure S5. Systemic delivery of miR-29 into mdx diaphragm. Table S1. List of upregulated genes after miR-29 injection. Table S2. List of downregulated genes after miR-29 injection. Table S3a. Functional annotation clustering of upregulated genes using DAVID GO. Table S3b. Functional annotation clustering of downregulated genes using DAVID GO. Materials and Methods.

Acknowledgments

We thank Dennis Y.M. Lo and Rossa W.K. Chiu for granting us the access to GAIIx sequencer and their generous support throughout the course of this study. The work described in this paper was substantially supported by three General Research Funds (GRFs) from the Research Grants Council (RGC) of the Hong Kong Special Administrative Region, China (CUHK476309 and 476310 to H.W. and 473211 to H.S.), two CUHK direct grants to H.W. (2041492 and 2041662) and a CUHK Direct Grant to H.S. (2041474). The authors declared no conflict of interest.

Supplementary Material

Figure S1.

Intramuscular injection of miR-29 oligos restores miR-29 expression in mdx limb muscles and improved regeneration.

Click here for additional data file. (1.9MB, tiff)
Figure S2.

RNA-seq reveals that miR-29 downregulates fibrotic genes in mdx muscles.

Click here for additional data file. (356.8KB, tiff)
Figure S3.

Co-staining of MyoD and α-SMA in mdx myoblasts.

Click here for additional data file. (315KB, tiff)
Figure S4.

miR-29 is downregulated in dystrophic muscles.

Click here for additional data file. (331.4KB, tiff)
Figure S5.

Systemic delivery of miR-29 into mdx diaphragm.

Click here for additional data file. (24.5MB, tiff)
Table S1.

List of upregulated genes after miR-29 injection.

Click here for additional data file. (86KB, xls)
Table S2.

List of downregulated genes after miR-29 injection.

Click here for additional data file. (307KB, xls)
Table S3a.

Functional annotation clustering of upregulated genes using DAVID GO.

Click here for additional data file. (29.5KB, xls)
Table S3b.

Functional annotation clustering of downregulated genes using DAVID GO.

Click here for additional data file. (110.5KB, xls)
Materials and Methods.
Click here for additional data file. (30.5KB, doc)

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1.

Intramuscular injection of miR-29 oligos restores miR-29 expression in mdx limb muscles and improved regeneration.

Click here for additional data file. (1.9MB, tiff)
Figure S2.

RNA-seq reveals that miR-29 downregulates fibrotic genes in mdx muscles.

Click here for additional data file. (356.8KB, tiff)
Figure S3.

Co-staining of MyoD and α-SMA in mdx myoblasts.

Click here for additional data file. (315KB, tiff)
Figure S4.

miR-29 is downregulated in dystrophic muscles.

Click here for additional data file. (331.4KB, tiff)
Figure S5.

Systemic delivery of miR-29 into mdx diaphragm.

Click here for additional data file. (24.5MB, tiff)
Table S1.

List of upregulated genes after miR-29 injection.

Click here for additional data file. (86KB, xls)
Table S2.

List of downregulated genes after miR-29 injection.

Click here for additional data file. (307KB, xls)
Table S3a.

Functional annotation clustering of upregulated genes using DAVID GO.

Click here for additional data file. (29.5KB, xls)
Table S3b.

Functional annotation clustering of downregulated genes using DAVID GO.

Click here for additional data file. (110.5KB, xls)
Materials and Methods.
Click here for additional data file. (30.5KB, doc)

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