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The complex process of skeletal muscle differentiation is organized by the myogenic regulatory factors (MRFs), Myf5, MyoD, Myf6, and myogenin, where myogenin plays a critical role in the regulation of the final stage of muscle differentiation. In an effort to investigate the role microRNAs (miRNAs) play in regulating myogenin, a bioinformatics approach was used and six miRNAs (miR-182, miR-186, miR-135, miR-491, miR-329, and miR-96) were predicted to bind the myogenin 3′-untranslated region (UTR). However, luciferase assays showed only miR-186 inhibited translation and 3′-UTR mutagenesis analysis confirmed this interaction was specific. Interestingly, the expression of miR-186 mirrored that of its host gene, ZRANB2, during development. Functional studies demonstrated that miR-186 overexpression inhibited the differentiation of C2C12 and primary muscle cells. Our findings therefore identify miR-186 as a novel regulator of myogenic differentiation.
Skeletal muscle differentiation (myogenesis) is a complex and tightly regulated process involving the commitment of embryonic precursors to the myogenic lineage, myoblast proliferation and exit from the cell cycle. In the final stage, mononucleated precursor cells fuse to form multinucleated mature myotubes (1,–4). This multistep process is orchestrated by a handful of genes coding the myogenic regulatory factors (MRFs).2 These MRFs belong to the basic-helix-loop-helix (bHLH) transcription factor family and consist of Myf5, MyoD, Myf6, and myogenin. Each MRF plays an important role in the process. Myf5 and MyoD are responsible for regulating the formation, proliferation, and longevity of myoblasts whereas Myf6 and myogenin play a critical role in the regulation of the final stage of differentiation (1, 5, 6).
Myogenin transcription is muscle specific and undergoes positive regulation both by itself and other helix-loop-helix proteins (7,–9). The myogenin gene is activated early on in development where it requires a change in DNA methylation status. In C2C12 myoblasts, this change can be seen as early as 2 h after induction of differentiation (10). Mutational analysis shows that myogenin transcription depends on two motifs, a consensus binding site for bHLH proteins and a consensus binding site for serum response factor proteins (11). Furthermore, MyoD and Myf5 are unable to substitute myogenin's functions during differentiation (12). Mice lacking the myogenin gene express normal levels of MyoD and Myf5 (13). Once activated, the expression of myogenin permits the differentiating myoblasts to undergo terminal myogenesis and fuse to form myofibres (14,–17). Myogenin's essentiality during differentiation is depicted in mice lacking the myogenin gene. The mice die soon after birth due to severe skeletal muscle deficiency, as myoblasts are unable to fuse into multinucleated myofibres (13, 18). Although the function of myogenin during development has proven to be essential, its role in adult myofibres is less defined (19). Interestingly, myogenin has been shown to serve as an important transcription factor regulating muscle metabolism and energy utilization in adult muscle (20). Myogenin is also expressed in regenerating adult myofibres where its expression is induced 4–5 days after muscle damage (21). Upon formation of mature myofibres, myogenin is eventually down-regulated (19, 22). We have recently shown that the down-regulation of endogenous myogenin gene expression reversed differentiation of C2C12 muscle cells (23). Moreover, in a conditional myogenin knock-out in regenerating muscle, its role appeared to be less critical (24). Finally, myogenin expression has been found to down-regulate genes involved in cell cycle progression leading to an inhibition of proliferation (25). The expression of MRFs must be precise both spatially and temporally. The level of MRF expression is controlled by the transcription of their corresponding genes and also by the stabilization of their mRNAs.
miRNAs are highly conserved, 22 nucleotide double-stranded RNA, which by binding the 3′-UTR of their target mature RNA (mRNAs) either mediate mRNA degradation or the inhibition of translation (26,–28). The 5′-end of the miRNA, primarily bases 2–8, is called the seed sequence and is the region which interacts with the seed matched region in the 3′-UTR region of its mRNA target (26). To date, 2555 unique mature human miRNAs have been identified (miRbase release 20), which are thought to play an important role in a wide range of biological processes, such as cell proliferation, stress response, differentiation, and apoptosis (29). The miRNAs involved in muscle differentiation fall into two categories, muscle specific and non-muscle specific. The most widely studied muscle-specific miRNAs are miR-1, miR-133, and miR-206. These miRNAs are regulated by the MRFs (30). Non-muscle specific miRNAs on the other hand, are expressed in muscle and other tissues (31, 32). These miRNAs mostly regulate muscle genes and transcription factors involved in myogenesis. For example, miR-31 targets Myf5, preventing Myf5 protein accumulation in quiescent satellite cells (33). miR-148 promotes myogenic differentiation by targeting the ROCK1 gene, a known inhibitor of myogenesis (34). miR-27 targets the paired box family of transcription factor 3 (PAX3) mRNA in skeletal muscle stem cells thus regulating the entry of cells into the myogenic program (35). Additionally, miR-27 has also been identified as a key regulator of Myostatin, a known inhibitor of myogenesis, which promotes myoblast proliferation (36). These data illustrate the importance of understanding how miRNAs control fundamental and complex biological processes such as myogenesis.
To date, the direct regulation of myogenin by miRNAs has not been demonstrated. Our investigation focused on the identification of miRNAs that could bind to and regulate myogenin. Our results show that miR-186 is a negative regulator of myogenin and inhibits myogenic differentiation.
C2C12 mouse myoblasts (ECACC) were grown to confluency under 5% CO2 at 37 °C in growth medium (GM), DMEM medium (Invitrogen) supplemented with 10% (v/v) fetal bovine serum (FBS) (Invitrogen), 2 mm glutamine (Invitrogen) and penicillin-streptomycin (100 mg/ml-100 units/ml) (Invitrogen). To differentiate, cells were then switched to differentiation medium (DM), DMEM supplemented with 2% horse serum (v/v) (Invitrogen), 2 mm glutamine, and penicillin-streptomycin (100 mg/ml-100 units/ml) for 4 days. HeLa cells were grown to 90% confluency before being subjected to transfections in GM as above. For primary mouse cell extraction, 8-week-old mice were killed by cervical dislocation, and the muscles were carefully removed from the hind limb. Myofibres were isolated from the extensor digitorum longus (EDL) muscle as described (37) and digested in 0.2% Collagenase Type 1 for 2 h at 37 °C. To purify muscle fibers from fibroblasts and cell debris, the digested muscle was washed by carefully pumping the muscle through pipettes of decreasing size and sequentially transferred to new, BSA-treated wells until the population of cells consisted purely of muscle fibers. The washed muscle fibers were next placed in Corning® 6-well plates coated with 1 mg/ml Matrigel (Collaborative Research) and incubated in activation media (DMEM with 10% (v/v) horse serum (Invitrogen), 0.5% (v/v) chick embryo extract (ICN Flow), 400 mm l-glutamine (Sigma), and 1% (v/v) penicillin/streptomycin solution (Sigma) at 37 °C for 48 h. The muscle fibers were removed from the plate by gentle blowing leaving the satellite cells attached to the plate's monolayer. The satellite cell population was further purified by a step involving re-plating of the remaining population into non treated plates, where the satellite cells remain in suspension. The suspended satellite cells were then plated into Corning® 6-well plates coated with 1 mg/ml Matrigel and maintained in activation media at 37 °C in 5% CO2.
a previously described muscle injury model was modified by injecting cardiotoxin (Sigma) intramuscularly into the tibialis anterior (TA) muscles of 8-week-old male C57BL/6 mice (38). The muscles were harvested at specific times (3, 5, 14 days) following injection. A saline-injected mouse was used as a negative control. The TA muscle was isolated and frozen, sectioned and stained for hematoxylin and eosin (H&E), or subjected to total RNA extraction.
100 pmol of each miRNA mimic, antimiR, negative miRNA, or anti-miR control were complexed with X-tremeGENE 9 transfection reagent (Roche) in Opti-MEM® reduced serum media (Invitrogen) before transfection. After 24 h, cells were then transfected with the wild type or mutated myogenin 3′-UTR luciferase plasmid (400 ng), and CMV-pRL Renilla (5 ng) (Promega, Madison, WI) plasmid in complex with Lipofectamine 2000 (Invitrogen). For transfections into C2C12 and primary mouse muscle cells, the same transfection procedure was repeated. For siRNA transfections in myotubes 100 pmol of siRNA myogenin (Invitrogen) were complexed with Lipofectamine RNAi/MAX (Invitrogen).
miRNAs were isolated using mirVanaTM miRNA Isolation Kit (Ambion) according to the manufacturer's instructions. Each sample (10 ng) was reverse transcribed into cDNA using the TaqMan® MicroRNA Reverse Transcription Kit (Applied Biosystems). Real-time PCR was performed in the Applied Biosystems 7900HT Fast Real-Time PCR System. miRNA expression was normalized to SNO135 (Applied Biosystems).
721 bp of the 753 bp long mouse myogenin 3′-UTR were amplified by PCR from genomic DNA extracted from the C2C12 cell line and then cloned into the multiple cloning site of the pMIR-Report luciferase miRNA expression reporter (Applied Biosystems catalogue no. AM5795). Similarly, 569 bp of the 779 bp long human myogenin 3′-UTR were amplified by PCR from human genomic DNA and cloned into the pMIR-Report luciferase miRNA expression reporter. 486 bp of the 2676 bp long mouse cyclin-D1 3′-UTR were directionally cloned into the same vector. The predicted miR-186 binding site (Fig. 1A) located in the mouse myogenin 3′-UTR was mutated by a substitution of 8 bp using the QuickChange Site Directed Mutagenesis Kit (Stratagene). All of the cloned vectors were verified by sequencing.
Genomic DNA was extracted from C2C12 myoblast using (QIAamp DNA Mini Kit, Qiagen) for sequencing of the miR-186 gene using the following primers: Intron 8 F 5′-CCGTGCTCACTTCAGACTGT-3′ and intron 8 R 5′-GACATCGCCCAGAAAAAGAA-3′. Total RNA was extracted from untransfected myoblasts or HeLa cells (Perfect RNAEukaryotic Mini kit, Eppendorf) and then subjected to reverse transcription. For semi-quantitative PCR, specific primers were used: Exon 7 F 5′-CAAGGTCTTCATCACGCTCA-3′ and exon 10 R 5′-GAACGGGAACCAGAATGTGT-3′. Luciferase F 5′-AATCTGACGCAGGCAGTTCT-3′ and Luciferase R 5′-CCAGGGATTTCAGTCGATGT-3′, GAPDH forward F 5′-TCATCATCTCCGCCCCTTCT-3′ and R 5′-GAGGGGCCATCCACAGTCTT-3′.
Following transfections, HeLa cells, C2C12 cells or primary mouse muscle cells were harvested, and assays were performed 24 h after the last transfection using the Dual-Luciferase Reporter Assay System (Promega, Madison, WI). Firefly luciferase activity was normalized to Renilla luciferase expression (internal control). Cells were subsequently lysed with commercial cell lysis buffer (Promega), and luciferase activity was measured using a luminometer (Berthold) according to kit protocols.
Proliferating myoblasts or differentiated muscle cells were subjected to protein extractions. 40–60 μg protein extracts were incubated with myogenin (1:200, abcam), ZRANB2 (1:400, abcam), cyclin-D1 (1:400, Santa Cruz Biotechnology), troponin (1:200, Santa Cruz Biotechnology), skeletal actin (1:200, Santa Cruz Biotechnology), Myf5 (1:400, Santa Cruz Biotechnology), MyoD (1:400, Santa Cruz Biotechnology), Myf6 (1:400, Santa Cruz Biotechnology) or GAPDH (1:1500, Santa Cruz Biotechnology) primary antibodies, followed by incubation with goat anti-mouse IgG or donkey anti-rabbit IgG secondary antibodies conjugated to horseradish peroxidase (Santa Cruz Biotechnology). Differentiated C2C12 myoblasts or primary mouse myoblasts were fixed in 4% paraformaldehyde and incubated with a monoclonal antibody against myosin heavy chain (MyHC) (Sigma) at a concentration of 1:400 in 1% BSA in PBS and a Texas-red-conjugated anti-mouse secondary antibody (Jackson Laboratories). Nuclei were stained with 4,6-diamidino-2-phenylindole (DAPI) (Vysis). Images were taken using a Zeiss Axiovision digital camera and then assembled using Adobe Photoshop Software. Cells were counted, at least three times from each pool of clones in ten different cellular areas to determine the fusion index (percentage of nuclei covered under myotubes over total nuclei).
ANOVA and Student's t test statistical tests were used to determine whether specific group mean differences were significant. The level of significance was set at 0.01. Data are presented as mean ± S.D.
To identify miRNAs involved in the regulation of myogenin, the 3′-UTR of mouse myogenin was screened for potential miRNA binding sites using miRanda software. By using certain criteria, such as conservation between species, co-expression in the same tissue and opposite expression profiles, the possible candidate miRNAs were narrowed down. Based on the miRSVR scoring, miR-182, miR-186, miR-135, miR-491, miR-329, and miR-96 were identified as candidates for binding myogenin. To investigate if the candidate miRNAs targeted the myogenin 3′-UTR, the entire wild-type mouse myogenin 3′-UTR was inserted downstream of the luciferase gene and assayed in HeLa cells (Fig. 1A). The mouse myogenin transcript was predicted to contain one canonical miRNA response element (MRE) for each of the candidate miRNAs. The initial screening of the candidate miRNAs indicated that miR-186 repressed luciferase activity most efficiently (~80%), compared with the remaining candidates (Fig. 1A). The binding site of miR-186 to the mouse myogenin 3′-UTR was predicted to occur toward the end of the 3′-UTR (Fig. 1B). The binding energy predicted for miR-186 was likeliest to bind the myogenin 3′-UTR and the binding site was conserved greater between closely related species compared with divergent species indicating possible evolutionary importance (Fig. 1C). The human myogenin 3′-UTR was assayed under the same conditions to investigate whether the slight change in the binding site MRE for miR-186 could cause repression of luciferase activity as with mouse myogenin 3′-UTR. The assay however revealed that the human myogenin 3′-UTR caused only a ~30% repression in two of the 6 miRNAs assayed, miR-182 and miR-186, implying that there is a possibility that the mechanism might be more efficient in mouse compared with its human counterpart (Fig. 1D). Introduction of mutations at the mouse seed matched region of miR-186 and transfections with a miR-186 mutant variant reverted the luciferase activity in HeLa back to original levels, indicating miRNA-mRNA specificity (Fig. 2, A and B). Luciferase 3′-UTR assays showed a similar result when performed in muscle cell environment and more specifically in C2C12 cells and mouse primary muscle cells (Fig. 2C). In the same experiments, antagomir-186 transfections caused an increase in the luciferase levels, indicating repression of endogenous miR-186 levels. Furthermore, to elucidate whether miR-186 exerts its effects at the protein or RNA level, the luciferase gene was PCR amplified following overexpression of miR-186 along with the mouse myogenin 3′-UTR luciferase construct into HeLa cells. Luciferase RNA expression levels fell more than 50% compared with the negative control (negative miRNA) indicating that miR-186 exerts its effects at the RNA level, which translates into the repression observed under the conditions of the luciferase reporter assay (Fig. 2D).
Following investigation of the genomic sequence of miR-186, it was identified that it is contained within the intron between exon 8 and 9 of the ZRANB2 gene (Fig. 3Ai). The position of the miR-186 gene was verified by DNA sequencing (data not shown). RNA analysis was performed and showed that the intron containing the miR-186 gene is spliced out of the ZRANB2 gene (Fig, 3Aii). This was also verified by cDNA sequencing (data not shown). A significant fraction of miRNA genes reside in the introns of host genes, in the same orientation and are thought to be co-expressed from the host gene's mRNAs (39,–41). We thus investigated the expression profile of miR-186 and its host gene, ZRANB2. The expression profile between the two molecules was similar during differentiation of muscle cells; they were both expressed at high levels before differentiation and gradually decreased toward the end of differentiation (Fig. 3, B and C).
Similarly, to associate the expression of miR-186 with the expression of myogenin during differentiation, myogenin expression levels were detected at various stages before and during differentiation. The expression profile of miR-186 shows an increase up until the first day of differentiation, after which it decreases gradually until the last day of differentiation where mature myotubes are formed. Meanwhile, myogenin protein levels were found to be minimal before being induced to differentiate, after which point, the protein levels gradually increased up until day 3 of differentiation, where the peak is reached. Myogenin protein levels then gradually declined in mature myotubes.
To investigate whether miR-186 mediates the post-transcriptional regulation of endogenous myogenin levels, mimics were introduced into C2C12 cells on the second day of differentiation, where endogenous miR-186 levels begin to decline. As a negative control, an RNA duplex of scrambled sequence was also transfected. Analysis of the protein levels showed that ectopic expression of miR-186 decreased endogenous levels of myogenin, indicating that miR-186 represses myogenin through its transcripts (Fig. 4A). As a next step, an antisense miRNA inhibitor was used to test whether the endogenously expressed miRNA has the ability to regulate myogenin levels. miR-186 inhibitor (amiR-186) was introduced on the first day of differentiation, where endogenous miR-186 expression is at its peak. Myogenin protein levels were increased when transfected with amiR-186, thus confirming the specific regulation of myogenin by miR-186 (Fig. 4A). The increase of myogenin protein levels, after amiR-186 transfection, raised the question of whether this could be the case, when the cells are at the proliferating stage, where myogenin and miR-186 are at low and high levels, respectively. To investigate this, C2C12 myoblasts were transfected during the proliferative stage with amiR-186, and myogenin protein levels were measured (Fig. 4A). There was no difference between transfected and control-transfected (anti-miR control) cells, indicating that reducing the endogenous miR-186 levels had no effect on the endogenous myogenin levels. It may well be possible that in proliferating muscle cells, myogenin levels are low due to its transcriptional regulation. This could mean that myogenin endogenous levels are so low that miR-186 cannot exert its effect on its transcripts. Additionally, it is possible that during proliferation of muscle cells, miR-186 is acting on a different target. To investigate the first scenario, and prove that miR-186 inactivity is due to the low myogenin levels, dose-response experiments were performed in proliferating myoblasts using the luciferase-3′-UTR-myogenin construct (Fig. 4B). Increased luciferase expression was responsive to miR-186 activity. More specifically, when transfected with minimal amounts of myogenin 3′-UTR plasmid, no difference was observed in luciferase activity between the amiR-186 and negative-transfected cells. As greater amounts of the luciferase plasmid were added, an increase in the luciferase values was observed compared with the negative control. Although the changes were not very high, albeit statistically significant, this result may hint that miR-186 activity depends on the endogenous levels of its target and has no effect at the proliferative stage, where myogenin levels are minimal (Fig. 4B).
To investigate the second scenario in which miR-186 is acting on a different target at the proliferative stage, a bioinformatics target search was carried out. Having gone through the possible candidates for miR-186, Cyclin-D1 came up as a possible candidate, which is a known regulator of cell cycle exit (42). We therefore investigated whether miR-186 could down-regulate mouse cyclin-D1 through its 3′-UTR. However, no significant repression was detected after transfection with miR-186 mimics, compared with the negative control (negative miRNA) (Fig. 4C). Similarly, no significant increase in cyclin-D1 was observed following transfection of amiR-186 in C2C12 myoblasts during the proliferative stage (Fig. 4C). It can be therefore concluded that cyclin-D1 is not an experimental target for miR-186 in these cells, and it is possible that the latter may be acting on other targets during proliferation of muscle cells.
Results presented so far indicate that miR-186 represses myogenin. As a final step, the ability of miR-186 to regulate muscle cell differentiation, through myogenin was investigated. C2C12 cells were transfected with miR-186 mimics and left to differentiate to form multinucleated myotubes. At the end of the process, cells were immunostained against MyHC, a late marker of differentiation (Fig. 5A). Muscle cells transfected with miR-186 mimic had lower ability to form myotubes, as compared with the negative controls (negative miRNA and mir-186 mutant) (Fig. 5A). The effect of miR-186 on differentiation, through myogenin repression was compared with a specific siRNA myogenin repression (Fig. 5A).
Based on the above results, it can be then concluded that by increasing miR-186, muscle differentiation is decreased, as expected from the decrease in myogenin protein levels. To confirm further the role of miR-186 on the process of muscle differentiation, amiR-186 was similarly transfected in C2C12 muscle cells. Reduction in the levels of endogenous miR-186 resulted in increased differentiation (Fig. 5A). To test whether the changes in differentiation were due to altered myogenin levels and not because of changes in the MRFs, MyoD, Myf5, and Myf6 protein levels were detected under the same conditions. The MRFs showed no significant changes compared with their respective negative controls. This indicates that the changes seen in muscle cell differentiation was a result of miR-186 targeting myogenin levels and not the other MRFs. Furthermore, to characterize further the molecular changes of differentiation the muscle markers troponin and skeletal actin were also measured along with myogenin at the protein level under similar conditions (Fig. 5C). Marked changes seen in both troponin and skeletal actin protein levels were compatible with the changes observed in the differentiation for muscle cells.
The effect of miR-186 on myogenin and subsequently on muscle cell differentiation was next examined in muscle cells, isolated directly from mouse. Importantly, we found that miR-186 mimic transfection in primary mouse cells displayed a decrease in myogenin protein levels (Fig. 6A). Similarly, amiR-186 transfection also caused a significant increase in myogenin protein levels, as was the case in the C2C12 cells (Fig. 6B). The effect of miR-186 on differentiation by transfecting with mimic or antimiR was then investigated in the primary muscle cells (Fig. 6C). By changing miR-186 levels, muscle cell differentiation in primary muscle cells was also changed. This proves that miR-186 is a potent inhibitor of differentiation and exerts its effect through down-regulation of myogenin.
To further explore the role of miR-186 in muscle differentiation, we investigated its role in muscle regeneration through a well-established mouse muscle injury model. The TA muscles of mice injected with cardiotoxin were harvested at specific days following injection (Fig. 7A). Maximum damage was observed 3 days after injections and almost complete recovery by day 14; myogenin RNA levels were found mostly increased during the stages of maximum damage, indicating induction of regeneration, similar to previous studies (38, 43) (Fig. 7B). The endogenous miR-186 levels were then measured using Real-Time PCR in order to follow the expression profile of miR-186 during skeletal muscle regeneration (Fig. 7C). They were found to be at very low levels during maximum muscle injury and when myogenin levels were high. The opposing endogenous expression levels of miR-186 and myogenin suggests a possible novel regulatory mechanism of muscle regeneration through miR-186. In conclusion, in this report we present for the first time evidence showing miRNA-mediated post-transcriptional regulation of myogenin, a major transcription factor in skeletal muscle.
The aim of this study was to identify miRNAs that regulate myogenin, a major transcription factor in muscles. Results from this study show that miRNA-186 is a novel post-transcriptional regulator of myogenin during skeletal myogenesis.
Although there is a great deal of information on the underlying mechanisms involved in the post transcriptional regulation of myogenesis, much more work remains to be done in the field of miRNA regulation of myogenesis (44). Of the six miRNAs tested, miR-186 showed a 5-fold decrease in relative luciferase expression compared with the negative miRNA, indicating that it is a strong candidate for binding to the mouse myogenin 3′-UTR. Additionally, conservation in the myogenin 3′-UTR between mouse, rat and human provides further evidence of possible function although the human myogenin 3′-UTR was not repressed to the same extent as its mouse counterpart.
Introducing the miR-186 and myogenin 3′-UTR into a controlled environment provides evidence that the two are able to interact when brought together. However, miRNA and target must be present in the same tissue if this is to be relevant in any way. Apart from scoring highly using target predicting software, miR-186 was chosen because of its expression during C2C12 muscle cell differentiation (30). This was further validated by RNA analysis.
The selection of miR-186 as the candidate for binding to myogenin 3′-UTR was finally confirmed by the following findings: (1) luciferase assays using wild type of mutant plasmids for the seed matched region of miR-186 in myogenin 3′-UTR along with a miR-186 mimic, demonstrated that only the co-transfection with the wild type reporter and the miR-186 mimic could dramatically reduce luciferase activity, (2) overexpression of miR-186 suppressed myogenin translation in C2C12 muscle cells and (3) inhibition of miR-186 in C2C12 muscle cells increased the translation of myogenin as was verified by protein analysis.
miR-186 resides between exon 8 and 9 of the ZRANB2 gene and it has been found that miRNAs are transcribed in parallel with their host transcripts (45). We therefore compared the protein levels of ZRANB2 with the expression levels of miR-186 during muscle differentiation and observed that the pattern was highly similar. Interestingly, by using primers annealing to exons on either side of the miR-186 gene, it can be concluded that the miR-186 is spliced out along with the entire intron to undergo further processing.
Although inhibition of miR-186 during the differentiating stage increased the levels of myogenin protein, this was not the case at the proliferating stage even though endogenous miR-186 levels were high. These and dose-response experiments showed that inhibition by miR-186 was not detectable when myogenin mRNA levels were low but is possible when the limiting factor is removed; i.e. the levels of myogenin were increased. Previous reports show that myogenin is down-regulated at the transcriptional level during proliferation of myoblasts, so miR-186 may be acting on a different target during this stage (19, 46, 47). Following screening for possible targets of miR-186 at the proliferative stage, we decided to select mouse cyclin-D1 as it is also expressed both in the pathway of interest and at the specific stage of interest. However, miR-186 did not inhibit cyclin-D1 at this stage suggesting other targets involved in regulation of the cell cycle of muscle cells remain to be identified. It is important to note however that a very recent report has shown that miR-186 targets cyclins, including cyclin-D1, which regulates the cell cycle in human cancer cell lines (48). Based on bioinformatics analysis, however, there is no conservation in the 3′-UTR of cyclin-D1 between mouse and human. It can be therefore concluded that cyclin-D1 is not an experimental target for miR-186 in these cells, and it is possible that the latter may be acting on other targets during proliferation of muscle cells.
To further elucidate the role of miR-186 in muscle differentiation, we next performed experiments to characterize the cellular effect that miR-186 had on muscle differentiation. Here, cells were stained against a marker of late differentiation, MyHC. The results were quantitated by measuring Fusion Index, which is the ratio of nuclei found within myotubes, identified as having 3 or more nuclei, against the total number of nuclei. These experiments were initially performed using the C2C12 cell line and then for further validation, repeated using primary mouse myoblasts.
Despite being expressed in skeletal muscle and cardiac muscle miR-186 is non-muscle specific and has been found to play a role in cell cycle regulation in cancer cells as well (30, 48,–50). It resides within an intron of the zinc finger, RAN-binding domain containing 2 (ZRANB2) gene, and we have shown that miR-186 expression mirrors that of its host gene ZRANB2. ZRANB2 is a spiceosomal protein involved in alternative splicing and is ubiquitously expressed in various tissues (51,–53).
Furthermore, a mouse muscle injury model was incorporated into this study in order to monitor the effects of alterations of endogenous miR-186 levels on muscle regeneration. The expression profile during mouse muscle regeneration of genes involved in both muscle differentiation and cell cycle regulation was studied. The opposing expression levels of miR-186 and myogenin at the chosen time points suggest a possible regulatory mechanism of muscle regeneration through miR-186.
Our study provides for the first time evidence for a miRNA-mediated mechanism of myogenin regulation. Myogenin is one of the four main MRF which contribute toward muscle formation and its transcriptional regulation is well known (19). miRNA-mediated post-transcriptional regulation by miR-186 and possibly other miRNAs provide an additional mechanism by which myogenin and other muscle-promoting molecules are regulated to maintain the balance between an active cycle and muscle cell differentiation.
*This work was supported by an A. G. Leventis Foundation grant and scholarship (to L. A. P. and A. A., respectively).
2The abbreviations used are: