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. Author manuscript; available in PMC: 2016 Feb 17.
Published in final edited form as: Cell Rep. 2015 Aug 28;12(10):1678–1690. doi: 10.1016/j.celrep.2015.07.066

TNFα-induced microRNAs control dystrophin expression in Becker muscular dystrophy

Alyson A Fiorillo 1, Christopher R Heier 1, James S Novak 1, Christopher B Tully 1, Kristy J Brown 1,2, Kitipong Uaesoontrachoon 1, Maria C Vila 1, Peter P Ngheim 1, Luca Bello 1,4, Joe N Kornegay 3, Corrado Angelini 4, Terence A Partridge 1,2, Kanneboyina Nagaraju 1,2, Eric P Hoffman 1,2,*
PMCID: PMC4757433  NIHMSID: NIHMS713383  PMID: 26321630

Summary

The amount and distribution of dystrophin protein in myofibers and muscle is highly variable in Becker muscular dystrophy and in exon-skipping trials for Duchenne muscular dystrophy. Here, we investigate a molecular basis for this variability. In muscle from Becker patients sharing the same exon 45–47 in-frame deletion, dystrophin levels negatively correlate with microRNAs predicted to target dystrophin. Seven miRNAs inhibit dystrophin expression in vitro, and three are validated in vivo (miR-146b/miR-374a/miR-31). microRNAs are expressed in dystrophic myofibers and increase with age and disease severity. In exon-skipping treated mdx mice, microRNAs are significantly higher in muscles with low dystrophin rescue. TNFα increases microRNA levels in vitro while NFκB inhibition blocks this in vitro and in vivo. Collectively, these data show that microRNAs contribute to variable dystrophin levels in muscular dystrophy. Our findings suggest a model where chronic inflammation in distinct microenvironments induces pathological microRNAs, initiating a self-sustaining feedback loop that exacerbates disease progression.

Graphical abstract

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Introduction

Duchenne muscular dystrophy (DMD) is caused by mutations in the dystrophin (DMD) gene that disrupt the open reading frame and prevent its protein translation (Chamberlain et al., 1987; Hoffman et al., 1987a; Hoffman et al., 1988; Hoffman et al., 1987b). Becker Muscular Dystrophy (BMD) is less severe and results from DMD mutations that preserve the reading frame. BMD-causing mutations lead to translation of a truncated dystrophin which is expressed at lower and more variable levels than full-length dystrophin (Beggs et al., 1991; Hoffman et al., 1989; Kesari et al., 2008).

Dystrophin content in BMD muscle varies within myofibers, between adjacent fibers, and between different patients, even when the same deletion mutation is shared. Dystrophin levels partly correlate with disease severity. Compared to normal muscle, dystrophin levels of ~3–15% are seen in severe BMD while >20% are associated with milder disease (Hoffman et al., 1988; Hoffman et al., 1989). BMD genotype-phenotype associations have previously been investigated to determine if there is a mutation-specific basis for inter-patient variation in dystrophin levels (Beggs et al., 1991; Cirak et al., 2011; Kesari et al., 2008; Koenig et al., 1987; Mendell et al., 2013; van den Bergen et al., 2014). These studies show that while greater disease severity is seen with amino-and carboxyl-terminal deletions, there is high variation in both dystrophin expression and clinical symptoms in patients with mutations in the central rod domain, even when the same exons are deleted.

The most common in-frame BMD deletion is of exons 45–47 (BMD Δ45–47) which codes for 150 amino acids in the central rod domain. We and others have reported variable dystrophin in BMD Δ45–47 muscle (5–80% (Kesari et al., 2008; van den Bergen et al., 2013)). These studies found little correlation between dystrophin amount and clinical phenotype, however BMD patients with <10% dystrophin exhibited a more severe clinical picture (Kesari et al., 2008; van den Bergen et al., 2013). BMD Δ45–47 patients should in theory, show similar gene expression, comparable mRNA stability, and produce an identical truncated protein with equivalent levels/stability. In contrast, the observed dystrophin content in these muscles varied significantly suggesting a mechanism of post-transcriptional dystrophin regulation.

A promising approach to induce de novo dystrophin in DMD muscle is exon skipping, where antisense oligonucleotides drive alternative splicing to produce a BMD-like dystrophin protein product. While extensive pre-clinical studies have provided proof-of-principle of this approach, dystrophin levels varied within and between muscle groups (Yokota et al., 2009; Yokota et al., 2012). Two clinical trials have also observed uneven dystrophin rescue (Cirak et al., 2011; Mendell et al., 2013).

We hypothesized that molecular mechanisms causing variable dystrophin protein levels in BMD are shared with those causing variability in exon skipping. To prevent introduction of confounding variables (differences in dystrophin transcript and protein stability), we utilized BMD muscles from patients with the same dystrophin Δ45–47 exon deletion as the initial discovery data set. Our preliminary data showed that dystrophin mRNA levels are maintained in BMD Δ45–47 muscle while dystrophin protein levels are variable. Given this, we investigated the role of microRNAs (miRNAs) in regulating post-transcriptional dystrophin levels.

Results

Variable dystrophin in Δ45–47 BMD patient muscles does not correlate with transcript levels

We carried out studies on 10 BMD patient biopsies harboring an exon 45–47 deletion mutation (BMD Δ45–47, Table S1). Dystrophin Western blot was performed with patient muscle and a standard curve of healthy muscle (‘Normal’) showing a dynamic linear range (Figure 1A, S1A). Normalized dystrophin was variable, ranging from 8%–63% (Figure 1B).

Figure 1. BMD Δ45–47 muscle shows variable dystrophin protein levels.

Figure 1

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(A) Western blot of BMD Δ45–47 muscle demonstrates variable dystrophin. Desmin and Coomassie-stain for myosin heavy chain (MYHC) used as loading controls. (B) Dystrophin transcript levels (RT-qPCR) do not correlate with dystrophin protein (Spearman’s correlation; rs=−0.0332; p= 0.412). (C–E) DTMs increase in BMD ‘low’ dystrophin muscle. TaqMan TLDA miRNA arrays performed with ‘Normal’ (n=6) and BMD Δ45–47 (n=10) muscle. (C) Elevated DTMs in BMD muscle with low dystrophin. Table lists DTMs elevated in BMD ‘High’ and ‘Low’ muscle; fold changes and p-values shown (‘High’ >20% dystrophin (n=6) and ‘Low’ <20% dystrophin (n=4) vs. Normal (ANOVA with post-hoc contrast groups: Normal vs. ’Low’; Normal vs. ‘High’). (D) Inverse correlation between dystrophin and DTMs (defined in each sample using ≥1.5-fold increase vs. Normal). Plot shows miRNAs detected in individual muscle vs. % dystrophin (Spearman’s correlation; rs=−0.726; ***p<0.0006). (E) DTMs inversely correlate with dystrophin protein. Plot shows miRNAs vs. % dystrophin protein in BMD and normal muscle (Spearman’s correlation rs=−0.691;**p=0.0013; rs =−0.610, **p=0.0048; rs=−0.732, **p=0.0011 for miR-146b-5p, miR-374a, and miR-382, respectively). Refer also to Figure S1; Table S1.

For subsequent studies, samples were stratified based on dystrophin levels. Groups were defined as ‘High’ corresponding to >20% dystrophin and ‘Low’ corresponding to <20% dystrophin (Figure S1B) based on reports showing dystrophin levels greater than 20% are needed to fully protect muscle fibers (van Putten et al., 2012). Dystrophin mRNA measured by RT-qPCR showed no correlation with dystrophin protein (Figure 1B, Figure S1C). Neither RT-PCR using primers flanking exons 44 and 48 nor RT-qPCR using custom probes against alternatively spliced transcripts showed evidence of alternative splicing (data not shown).

Predicted miRNA binding sites in the dystrophin 3′UTR correspond to miRNAs elevated in dystrophic muscle

Given the lack of correlation between dystrophin mRNA and protein levels, we hypothesized miRNAs may post-transcriptionally regulate dystrophin. Seventy-eight potential miRNA binding sites for 67 distinct miRNAs were identified in evolutionarily conserved regions of the 2.7 kb dystrophin 3′ untranslated region (UTR) (Figure S1D). miRNA profiling was performed using Δ45–47 BMD (n=10), Normal (n=6), and DMD muscle biopsies (n=5) with TaqMan TLDA Arrays containing probes for 51/67 miRNAs predicted to bind the dystrophin 3′UTR. In BMD ‘Low’ samples, 14 miRNAs showed significant upregulation (1.5-fold to 17-fold, Figure 1C). In contrast only 5 miRNAs were elevated in BMD ‘High’ samples (Figure 1C). In an additional analysis, the number of elevated miRNAs (≤1.5 fold) in each BMD sample showed a modest inverse correlation when plotted as a continuous variable against dystrophin protein (Figure 1D). Similarly, individual miRNA levels (miR-146b, miR-374a and miR-382 shown as examples, Figure 1E) showed inverse correlations with dystrophin. Together, these data show dystrophin-targeting miRNAs (herein, referred to as DTMs) are inversely related to dystrophin levels in BMD Δ45–47 muscle.

In an additional analysis, we found five DTMs elevated (3.6 to 25.1-fold) in both DMD and BMD ‘low’ muscle (miR-146–5p, miR-382, miR-758, miR-214, and miR-494) (Figure S1E). Dystrophin-targeting miR-31 was also upregulated in DMD muscle (Figure S1E).

DTMs inhibit dystrophin protein translation in vitro

To determine if DTMs modulate dystrophin protein levels, we constructed a reporter containing the 2.7 kb dystrophin 3′UTR downstream of Renilla reniformis luciferase; this reporter co-expressed Firefly luciferase from a separate promoter, thus providing a robust internal transfection control (Figure 2A). This reporter was co-transfected into C2C12 myoblasts along with one of 14 miRNAs upregulated in BMD ‘Low’ muscle (Table S2). Seven miRNAs inhibited dystrophin expression (miR-31, miR-146a, miR-146b-5p, miR-223, miR-320a, miR-374a, and miR-382), two enhanced expression (miR-195/miR-758) and five had no effect (Figure 2B).

Figure 2. miRNAs inhibit dystrophin protein translation in vitro.

Figure 2

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(A) Schematic of dystrophin 3′ untranslated region (UTR) reporter. The human dystrophin 3′UTR was cloned into the 3′ end of a Renilla reporter gene (psi-CHECK2 vector). The psi-CHECK2 vector co-expresses Firefly luciferase, and thus provided an internal transfection control. (B) miRNAs inhibit dystrophin 3′ UTR reporter activity. Individually, 14 dystrophin mRNA-targeting miRNAs were co-transfected with reporter into cells; % inhibition is provided in graph (n=4 replicates; ANOVA**p<0.01; ***p<0.001; ****p<0.0001 vs. negative (−) control). (C) Western blot of healthy human myotubes transfected with 50nM of indicated miRNAs. Tubulin (loading control) and densitometry values (% CTRL) are provided. (D) DTMs show synergistic inhibition. The 3 most potent DTMs (1nM, miR-146b, miR-374a, and miR-31) were transfected into cells individually, or in combination (referred to as ‘Biomix’); results reported as % inhibition (n=4 replicates; ANOVA, **p<0.01; ***p<0.001; ****p<0.0001 versus negative (−) control). (E) Schematic shows base-pairing of miRNAs with dystrophin 3′ UTR; called miRNA recognition elements or MREs. MRE mutants were constructed as shown; 4–5 nucleotide substitutions were made to reporter (mutated nucleotides in red). For miR-146a/b sequence x=c, y=a for miR-146b and x=t, y=g for miR-146a (blue). Mutagenesis was performed on 1 of 3 miR-374a MREs, however this mutant was anticipated to have little effect on reporter expression due to 2 non-mutated miR-347a MREs remaining (gray). (F) MRE mutagenesis reduces dystrophin inhibition. 50nM indicated miRNAs were co-transfected into cells along with dystrophin wild-type (white bars) or a MRE-mutant 3′UTR reporter (black bars). Mutated MRE construct matches transfected miRNA for each condition as indicated (n=4 replicates; Student’s t-test for wild-type versus mutant; #p<0.1; *p<0.05; **p<0.01). Refer also to Table S2.

We tested the most potent DTMs in human myotubes in vitro. Immortalized human myoblasts were transfected with the indicated miRNAs or control and differentiated into myotubes. Western blot showed all miRNAs reduced dystrophin to <20% of normal levels (Figure 2C).

Using miRNAs showing the strongest inhibition (miR-146b, miR-31, miR-374a), we determined if miRNAs could have an additive or synergistic effect in combination. Here, we created a miRNA mix (“Biomix”) composed of miRNA levels that approximated expression in dystrophic muscle biopsies based on Cq levels from miRNA arrays (70% miR-146b, 25% miR-374a, and 5% miR-31, Figure 2D, Table S2). At 1nM, the “Biomix” inhibited reporter activity more than any single miRNA at the same concentration (Figure 2D), indicating DTMs may work in concert to inhibit dystrophin.

To determine the specificity of miRNA-mediated dystrophin inhibition, we mutated specific miRNA response elements (MREs) in the 3′UTR reporter (Figure 2E). Mutants were made for each miRNA in the Biomix including disruption of one miR-374a MREs. We anticipated, however, this mutant would have little or no effect due to the other two MREs located within the dystrophin 3′ UTR. Results showed miR-146a/b MRE mutagenesis attenuated inhibition both of miR-146b and miR-146a and miR-31 MRE mutagenesis alleviated miR-31-specific inhibition (Figure 2F). miR-374a MRE mutagenesis, however, had no effect on miR-374a-mediated inhibition (Figure 2F), likely due to the two functional miR-374a MREs remaining.

DTMs regulate dystrophin in vivo

We next tested the effects of DTMs in wild-type mice in vivo. The Biomix (miRNA) was injected into the right tibialis anterior (TA) of 6-week-old C57BL10/J mice; the left TA received scrambled control (CTRL) (Table S2). Seven days post-injection muscles were harvested and analyzed for miRNA and dystrophin. RT-qPCR showed successful intramuscular delivery of exogenous miRNAs (Figure S2A). At the injection site (indicated by tattoo dye), immunofluorescence showed reduced dystrophin in miRNA, but not in CTRL-injected mice (Figure 3A). RT-qPCR showed dystrophin mRNA was not affected (Figure S2B), consistent with translation inhibition as the primary mechanism-of-action.

Figure 3. DTMs reduce dystrophin in vivo.

Figure 3

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(A–B). miRNA pool (called “Biomix” with 70% or 1.05μg miR-146b; 25% or 0.375 μg miR-374a; and 5% or 0.075 μg miR-31) was injected into TA of 6-week old C57BL10/J mice (group termed miRNA, n=6/group). Equivalent amount of control was injected into left TA (group termed CTRL). Muscles were harvested 7 days later. (A) miRNA injection of wild-type mice to observe effects on steady-state dystrophin. Left; Representative immunofluorescence images overlaid with tattoo dye from brightfield to delineate site of injection; red=dystrophin; green=tattoo dye; white arrows denote where dystrophin levels are decreased in miRNA, but not in CTRL injected muscles; scale bars=100μM. Right; Average pixel count (dystrophin levels) around injection site (Student’s t-test; *p<0.05). (B–C) miRNA injection after injury in wild-type mice to observe effects of de novo dystrophin expression. Muscle injury was inflicted using notexin. Three days post-injection miRNAs were injected into the right TA; CTRL was injected into the left. Mice were sacrificed 7 days post-injury (n=3/group). (B) miRNAs reduce dystrophin expression post-injury. Western blot of CTRL or miRNA-injected muscle to show dystrophin. Loading controls are provided. (C) Left; Dystrophin immunostaining in CTRL and miRNA-injected mice. Central nucleation demarcates regenerated fibers (red=dystrophin, blue=DAPI). Right; average pixel count (dystrophin levels) per field (Student’s t-test; **p<0.01). Refer also to Figure S2, Table S2.

In the previous experiment, miRNA delivery was restricted to a small region surrounding the injection site. Thus, we performed a second experiment where muscles were injured via notexin prior to miRNA injection. This approach also had the effect of removing endogenous dystrophin in mature myofibers, enabling us to determine if injected miRNAs inhibit de novo dystrophin during myogenic regeneration (half-life of dystrophin in mature myofibers is ~2 months, (Wu et al., 2012)). The TA muscles of 6-week-old C57BL6 wild-type mice were injected with 100μg notexin. Four days after the initial injury, mice were injected with the miRNA Biomix or CTRL. Mice were harvested 7 days post-injury, when dystrophin expression becomes visible in myotubes (Hoshino et al., 2002). Western blot showed reduced dystrophin in miRNA but not in CTRL-injected mice (Figure 3B). Immunofluorescence corroborated these results (Figure 3C).

DTMs in dystrophic muscle increase with age

Golden retriever muscle dystrophy dogs (GRMD) exhibit variable histopathology, similar to DMD boys (Cooper et al., 1988; Shimatsu et al., 2005; Smith et al., 2011; Valentine et al., 1988). Given this, we assessed DTMs in GRMD muscle. We first measured DTMs in 6-month old GRMD vastus lateralis (VL) as it is histologically severe and similar to the VL of DMD patients. miR-146b, miR-146a, and miR-223 were elevated in the GRMD VL (n=9; 50-fold, 3-fold, and 8-fold, respectively) as compared to wild-type (Figure 4A, Table S3 for TaqMan assay information). DTMs were also measured in GRMD cranial sartorius (CS), which is mildly affected in both dogs and humans (Calabia-Linares et al., 2011; Lemaire et al., 1988; Nghiem et al., 2013). miR-146b, miR-146a, miR-223 and miR-382 were elevated (n=9; 25-fold, 1.3-fold, 2.5-fold, and 4-fold) in GRMD CS, albeit at slightly lower levels as compared to the GRMD VL (Figure S3A). Supporting this, DTMs were similarly elevated in mdx gastrocnemius (Figure S3B).

Figure 4. DTMs are elevated in dystrophic muscle and increase with age.

Figure 4

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(A) miRNAs are elevated in dystrophic dogs. Levels of miR-146b, miR-146a, miR-223 in the vastus lateralis (VL) muscle of 6-month old GRMD (n=9) compared to aged matched wild-type dogs (n=3) (Student’s one tailed t-test; #p<0.1, p<0.05, **p<0.01, ***p<0.001). (B) DTMs increase with disease progression. Levels of miR-146a, miR-146b, miR-223 in VL muscle biopsies of 1 and 6 month-old GRMD dogs (n=6/group). (C) DTMs increase with age in mdx mice. Left; DTMs in TA of mdx mice (12 days, n=3; 8 weeks, n=4). miR-223 and miR-31 levels are shown (Student’s t-test; ***p<0.001; *p≤0.05). Right; Representative H&E of cross-sections from TAs of 12-day and 8-week old mdx mice where scale bar = 100μM. (D–E) DTMs are elevated in whole extensor digitorum longus muscle (EDL) and in purified myofibers from mdx EDL. (C) DTMs in whole EDL of mdx or age-matched wild-type mice (Student’s t-test, n=4 per group; **p<0.01; *p<0.05; #p<0.1) (D) DTMs in purified myofibers from the contralateral EDL of the same mdx and wild-type mice from panel C. Note in C and D, miRNA upregulation is maintained in purified (n=4/group; Student’s t-test with Mann-Whitney correction for non-Gaussian distribution; **p<0.01; *p<0.05; #p<0.1). Refer also to Figure S3, Table S3.

We next determined if DTMs correspond with dystrophic disease severity, utilizing VLs from 1 or 6-month old GRMD and wild-type dogs. Three miRNAs (miR-146b/miR-146a/miR-223) increased with age (n=6; Figure 4B). Separately, we performed a smaller longitudinal analysis using VL muscle taken serially from GRMD (n=4) or wild-type (n=4) dogs over a period of 6 months. miR-146b, miR-146a and miR-223 increased in 100% of GRMD dogs from 1 to 6 months while in WT, these miRNAs either decreased or showed a smaller change (Figure S3C). To further demonstrate that DTMs are associated with disease progression, we analyzed TAs of 12-day and 8-week old mdx mice. We observed marked increases in miR-223 and miR-31 correlating with age (Figure 4C). miR-146a/miR-146b were equally elevated in both ages of mdx mice, but this likely due to early pre-symptomatic activation of NFκB as previously observed in newborn DMD patients (Chen et al., 2005). Consistent with previous studies (Hamrick et al., 2010), when DTMs were assessed in wild-type mice, only miR-382 levels were significantly altered (decreased to ~10% in older mice); however the change was opposite to the direction of the mdx genotype effect (Figure S3D).

To see if DTMs were expressed in muscle cells, mature myofibers were enzymatically dissociated from the extensor digitorum longus muscle (EDL) of 8-week-old mdx and wild-type mice. miR-146b, miR-146a, miR-31 and miR-223 were elevated in mdx whole EDL (Figure 4D) and in purified fibers (Figure 4E). In mdx myotubes in vitro, four DTMs increased during differentiation (Figure S3E), suggesting a role in regeneration. Together, these data show miRNAs become increasingly elevated in older dystrophic muscle, and DTMs are not associated with the normal aging process in muscle.

DTMs reduce exon skipping success

We tested whether DTMs contribute to variable dystrophin rescue observed in exon skipping. Four-week-old mdx mice were given a single 800 mg/kg intravenous injection of exon 23-targeting morpholino, and were sacrificed after 1 month. From one mouse, adjacent sections of TA, gastrocnemius, and diaphragm muscles were analyzed for dystrophin protein by mass spectrometry (Brown et al., 2013) and for DTMs by RT-qPCR. We detected high dystrophin levels in the TA (~40%) while diaphragm and gastrocnemius showed lower levels (5%, Figure 5A). Conversely, DTMs were low in the TA (miR-146a/miR-374a/miR-223/miR-320a/miR-382) and high in the gastrocnemius and diaphragm muscles (Figure 5A).

Figure 5. DTMs are inversely correlated with exon skipping success in vivo.

Figure 5

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(A–B) 4-week-old mdx mice were given PMO (single high intravenous dose, 800 mg/kg) driving exon 23 skipping. 4 weeks post-treatment, muscles were analyzed for miRNA expression via RT-qPCR and for dystrophin via SILAM Mass Spectrometry (n=3 muscles). (A) miRNAs influence intra-variability in dystrophin rescue. Dystrophin and miRNA levels are shown for tibialis anterior (TA), gastrocnemius (gastroc) and diaphragm muscle from a single PMO-treated mouse (ANOVA, *p<0.05; **p<0.01; ***p<0.001). (B) Inter-subject variability in dystrophin rescue influenced by miRNAs. Plot of dystrophin protein as % wild-type (y axis) and a combinatorial score of 7 miRNAs (miR-146b, miR-374a, miR-31, miR-223, miR-146a, miR-382 and miR-320a scored as low, moderate or high, x axis; 63 total measures). Measurements determined using triceps, TA and gastroc of treated mdx (ANOVA, **p<0.01).

To determine if DTMs contributed to intra-animal exon skipping variability, two additional morpholino-injected mice were studied. DTMs in mdx triceps, gastrocnemius and TA were measured (n=9 muscles tested; 7 miRNA measures/muscle). Muscles were stratified into miRNA ‘low’, ‘mid’, or ‘high’ groups and plotted against % dystrophin. We found the miRNA ‘low’ group showed quite high dystrophin while ‘high’ group had lower dystrophin levels (Figure 5B). These data suggest DTMs may contribute to inter and intra-subject variability of dystrophin rescue in exon skipping studies.

DTMs are induced by pro-inflammatory TNFα in myogenic cells

Two DTMs are induced by NFκB; miR-146a in THP-1 cells (Taganov et al., 2006) and miR-223 in T-cells (Kumar et al., 2014). We determined if DTMs could be induced by pro-inflammatory stimuli in mdx H2K myotubes. TNFα treatment increased miR-146a and miR-223 (Figure 6A) while pre-treatment with NFκB –inhibiting anti-inflammatory drugs (prednisolone, VBP15 (Heier et al., 2013)) suppressed induction (Figure 6A). Both drugs also decreased miR-146b and miR-382 levels, with VBP15 showing greater effects (Figure 6A).

Figure 6. DTMs are induced by NFκB-mediated inflammation.

Figure 6

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(A) mdx H-2K myotubes treated with indicated drug, were induced with TNFα; DTMs assayed by RT-qPCR. miR-146a and miR-223 increase with TNFα; VBP15 or prednisolone (Pred) pre-treatment inhibits induction; miR-146b and miR-382 decreased with VBP15, but not Pred (n=5/group; ANOVA, ****p<0.0001, ***p<0.001, **p<0.01; *p<0.05; #p<0.1). (B) Muscles from 6-month-old mdx mice treated with Pred (5 mg/kg/day) or VBP15 (45 mg/kg/day) as described (Heier et al., 2013). miR-146a, miR-146b and miR-223 decreased with both drugs (n=8/group; ANOVA; ****p<0.0001, **p<0.01, *p<0.05). (C) Muscles from 8-week-old mdx mice treated with Pred (5 mg/kg/day) or VBP15 (15 mg/kg/day) as described (Heier et al., 2013). miR-146a and miR-223 are reduced by both drugs, whereas miR-382 increases with Pred, but not VBP15 (n=5/group; ANOVA; **p<0.01, *p<0.05). (D) DTMs associated with the NFκB pathway are preferentially elevated in “Severe” BMD muscle. Left; Representative H&E staining showing “Mild” and “Severe” BMD pathology. Right; miR-146a, miR-146b and miR-223 levels in BMD “Mild and “Severe” muscle (Student’s t-test **p<0.01; *p<0.05; #p<0.1). Scale bars=200μM. Refer also to Table S4, Figure S4.

To investigate anti-inflammatory effects in vivo, miRNA levels were measured in archival samples from prednisolone or VBP15-treated mdx mice. Mice were 6-month-old mdx or age-matched wild-type treated for 4 months (Heier et al., 2013). Both drugs significantly reduced diaphragm miR-146a and 146b levels and slightly reduced miR-223 (Figure 6B).

Archival samples from gastrocnemius of 8-week old prednisolone or VBP15-treated mdx mice were additionally obtained from a separate study (Heier et al., 2013). Here, both prednisone and VBP15 reduced miR-146a and miR-223 in comparison to untreated mice while miR-382 was increased in prednisolone but not in VBP15-treated mice (Figure 6C). We also assessed BMD muscle biopsies (VL, n=9) with “Mild” or “Severe” histopathology (Table S4, Figure 6D). In this cohort, 5 of 7 DTMs were elevated as compared to healthy muscle (Figure S4). Interestingly, only those DTMs associated with the NFκB pathway (miR-146a, miR-223, miR-382) were elevated in “Severe” vs. “Mild” BMD muscle (Figure 6D). Collectively, these data show NFκB regulates a subset of DTMs and its inhibition reduces pathological miRNAs in muscle.

Discussion

Here, we utilize muscle from BMD patients harboring an exon 45–47 to model dystrophin protein variability observed in exon skipping studies. This enabled us to examine dystrophin regulation without confounding variables, such as protein stability, attributed to a specific DMD exon deletion (van den Bergen et al., 2014). Using this method, we provide insight into the molecular mechanisms contributing to variable dystrophin. We identify miRNAs that regulate dystrophin and are induced by inflammation, a feature of dystrophic muscle. Our findings suggest a model for dystrophin variability in muscle and for variable clinical progression of BMD patients sharing the exon deletion (refer to Graphical Abstract). As dystrophic myofibers remodel, they induce a pro-inflammatory response in distinct microenvironments, triggering immune cells to release inflammatory cytokines, such as TNFα. This activates NFκB signaling in myofibers, which induces DTM transcription (miR-146a, miR-223) which, in turn, inhibits dystrophin translation. These events could further exacerbate aberrant signaling that occurs in dystrophic myofibers and initiate a positive feedback loop which would 1) lead to further increases in DTMs and 2) would result in decreased, yet variable, dystrophin in individual fibers and muscle groups. Chronic activation of these processes would result in variable clinical phenotypes that would presumably worsen with age and disease progression. Inhibition of DTMs could theoretically increase dystrophin in BMD, and thus provides a potential therapeutic target.

Our findings contribute to the knowledge initiated by a few key bodies of work. One report showed miR-31 is associated with muscle regeneration and miR-223 is associated with inflammatory infiltration following muscle injury (Greco et al., 2009); another showed proof-of-principle that miR-31 can inhibit dystrophin in vitro (Cacchiarelli et al., 2011). Furthermore, previous studies of serum miRNAs show the extent of miRNA dysregulation is linked to age and a disease progression (Jeanson-Leh et al., 2014; Vignier et al., 2013)

DMD muscle shows variable histopathology that is, in part, due to asynchronous regeneration (Dadgar et al., 2014) which creates muscle microenvironments with various degrees of pro-inflammatory and pro-fibrotic networks. Our proposed model suggests inflammatory microenvironments influence dystrophin via DTM induction. Supporting this, previous reports show only a fraction of healthy donor myoblasts or bone marrow-derived cells produce dystrophin in DMD muscle, perhaps due to DTMs within the muscle microenvironment (Gussoni et al., 1997; Wernig et al., 2005). Additionally, here we show an inverse relationship between DTMs and dystrophin rescue in exon-skipping treated mdx. Given this, DTM inhibition may improve exon skipping success. Evidence for this includes an in vitro study where co-transfection of an exon-skipping lentiviral construct (U1#51) and a locked nucleic acid (LNA) to inhibit miR-31 resulted in increased dystrophin (Cacchiarelli et al., 2011).

Previously, our lab investigated mosaic female DMD carriers with different proportions of non-mutated dystrophin (Pegoraro et al., 1995). In these patients, some dystrophin-competent myonuclei failed to make dystrophin, specifically in older patients. Here, we show DTMs increase with age in GRMD and mdx muscle. This suggests age-related increases in miRNAs may contribute to the previously described failure of dystrophin-competent nuclei to produce dystrophin.

miRNA profiling in the aging heart has identified 65 miRNA that are differentially expressed (Zhang et al., 2012). This list included increases in miR-146a, miR-146b, miR-223 and miR-374a, which we have reported here. A separate study showed found the aging heart had reduced dystrophin levels (Townsend et al., 2011), suggesting a link between miRNAs and decreased cardiac dystrophin during aging.

Most miRNAs described here do not exhibit distinct tissue specificity. miR-31 is higher in normal human GI and epithelial tissues (Liang et al., 2007) and miR-146 is elevated in the murine heart (Lagos-Quintana et al., 2002). Other reports detect DTMs in skeletal muscle at different phases of muscle regeneration (Greco et al., 2009). We found elevated DTMs in purified mdx myofibers, suggesting muscle-specific miRNA expression. We also show TNFα induces miR-146a and miR-223 in mdx myotubes, while NFκB inhibition attenuates induction. While DTMs described here are not classically defined as ‘myomiRs’, previous reports show non-muscle specific miRNAs are also imperative for proper muscle function (Novak et al., 2013). One report shows miR-146b, miR-31, miR-223, known as ‘dystromiRs’ are differentially expressed in dystrophic muscle and have been shown to play a role in myogenesis and muscle regeneration (Roberts et al., 2012). Other reports show miR-146b-5p promotes myogenic differentiation (Khanna et al., 2014; Kuang et al., 2009) and miR-31 and miR-223 are induced in ischemia-damaged myofibers (Greco et al., 2009). Supporting thus, here we show DTM induction during myoblast differentiation. Together, these data suggest DTMs play a role in normal muscle regeneration.

Inflammatory cells could contribute to DTM induction in dystrophic muscle. In this scenario, “crosstalk” between immune cells and myofibers could be mediated by horizontal transfer of miRNAs by exosomes or microvesicles (Ismail et al., 2013; Kosaka et al., 2010; Mittelbrunn et al., 2011; Skog et al., 2008; Valadi et al., 2007; Zhang et al., 2010). While there are knowledge gaps in the mechanism of RNA transfer from immune to other cells, it is plausible that miRNAs could behave similar to endocrine peptide hormones where distant cytokines induce a positive autocrine/paracrine feedback loop (Clevenger and Plank, 1997).

We show TNFα-mediated NFκB activation induces DTM expression in dystrophic myotubes. This supports previous studies that identified NFκB consensus elements in miR-146a/miR-223 promoters (Kumar et al., 2014; Taganov et al., 2006). In treating DMD, patients undergoing exon skipping therapy will likely be co-administered glucocorticoids. One inclusion criteria for recent exon skipping trials was >24 weeks of glucocorticoid treatment (Mendell et al., 2013). Glucocorticoids globally effect inflammation and gene transcription, which could impact DTM expression. Here, we show prednisolone and VBP15 reduce DTMs miR-146a, miR-146b and miR-223, suggesting these drugs could increase dystrophin if combined with exon skipping. Prednisolone also increased miR-382, whereas VBP15 had no effect. This difference could be explained by the ability of VBP15 to dissociate glucocorticoid receptor-mediated transactivation activity. These data suggest anti-inflammatory compounds such as prednisolone and particularly VBP15 could enhance exon skipping success (Heier et al., 2013).

It is conceivable that DTMs are involved in other muscle disorders where NFκB signaling is enhanced. Interestingly, a previous report showed elevated DMTs in a wide variety of muscle disorders such as Myositis, Myoshi myopathy and Limb Girdle Muscular Dystrophy (Eisenberg et al., 2007). Thus, DTMs may be a common signature of muscle diseases where chronic inflammation is present and could potentially provide therapeutic targets for a broader range of muscle disorders.

Consistent with our findings, a previous study showed that miR-31 represses dystrophin through the 3′ UTR (Cacchiarelli et al., 2011), results that were validated here. In dystrophic muscles, we detected miR-31 at lower absolute levels than other DTMs. miR-31 did not increase in GRMD muscle, and was not associated with exon skipping success mdx. Thus, while miR-31 was one of seven DTMs characterized, others appeared more relevant to both disease progression and therapeutics.

Our work here elicits questions regarding the role of the dystrophin 3′UTR in normal muscle, given the abundance of miRNA binding sites and the high conservation of this region. One model revolves around remodeling of myofibers in healthy muscle. Myofibers are one of the more morphologically adaptable cells and activity can result in rapid cell hypertrophy or atrophy. The normal function of dystrophin is to provide a rigid membrane cytoskeleton and robust connections between intracellular contracting myofibrils and the extracellular matrix. However, myofibers need to transiently destabilize the membrane cytoskeleton to remodel (Kee et al., 2004). Given previous reports showing that lengthening muscle contractions result in reduced dystrophin (Komulainen et al., 1998), it is possible that the “normal” role of DTMs is to enable transient dystrophin reduction in remodeling myofibers.

Here, we show proof-of-principle that dystrophin is reduced by inflammation-induced miRNAs that are elevated in dystrophic muscle. Our data provide insight into phenotypic discrepancies observed in BMD and variable success observed in DMD exon skipping clinical trials. We show NFκB inhibition, in addition to quelling inflammation, may provide the added benefit of increasing de novo dystrophin production. This work could potentially provide an avenue for molecular-based therapy for BMD patients and an adjuvant therapy in DMD to increase exon-skipping effectiveness.

Experimental Procedures

Muscle biopsies

All details are provided in Extended Experimental Procedures.

Western blot analysis

Muscle protein was extracted from cryosections with lysis buffer containing 75 mM Tris-HCl (pH 6.8), 10% sodium dodecyl sulfate, 10 mM EDTA, and 5% 2-mercaptoethanol as described (Yokota et al., 2009). Additional detail is in Extended Experimental Procedures.

TaqMan ® miRNA Low Density Arrays (TLDA)

RNA was extracted from 20 mg muscle (10 BMD; 5 DMD; 6 control) with TRIzol (Life Technologies). miRNA array was performed with TaqMan low-density Array A (TaqMan® Array Human miRNA, v3.0A; 382 miRNAs, Applied Biosystems, Life Technologies). Single-stranded cDNA was synthesized from 100 ng RNA using TaqMan® MiRNA Reverse Transcription Kit (Life Technologies) and RNA-specific stem-looped Megaplex RT Primers, Human Pool A v2.1 (Life Technologies). Additional details in Extended Experimental Procedures.

RT-qPCR assays

mRNA

Total RNA was extracted from muscle biopsies (~20 mg) using TRIzol (Life Technologies) according to manufacturer’s instructions with isopropanol precipitation performed at −20°C overnight. Total RNA was reverse-transcribed to cDNA using Qscript cDNA synthesis kit (Quanta) and then analyzed using human-specific TaqMan probes (Life Technologies) and the 7900HT Fast Real-Time PCR system. For TaqMan gene expression assay IDs, refer to Table S3. Muscle-specific gene expression was normalized to titin (TTN); whereas, inflammatory cell identifying transcripts were normalized to hypoxanthine phosphoribosyl transferase (HRPT). Results were calculated using the 2−ΔΔCq method (Livak and Schmittgen, 2001). Mouse gene expression was quantified in the same manner as above, with mouse-specific TaqMan probes (Life Technologies, refer to Table S3).

miRNA

Human miRNAs were quantified using TLDA Assays as above from RNA extracted from ~20 mg of muscle. Mouse and dog miRNAs were quantified (20 mg muscle) using TaqMan assays (Life Technologies) according to manufacturer’s protocol. Table S4 lists all miRNA Assay IDs.

Luciferase Assays

C2C12 myoblasts (3′UTR assay) or HEK 293 cells (mutagenesis assay) were seeded in 24-well plates at a density of 4×104 or 8×104 cells/well and co-transfected 24h later with 200 ng dystrophin 3′UTR or mutant reporter and with 50 nM miRNA mimics (Life Technologies, refer to Table S5) with lipofectamine 2000. Cells were harvested 24h later according to Dual-Glow Luciferase Reporter Assay System protocol (Promega). Results were normalized to an internal control driven by a separate promoter in the same reporter. Results are reported as % change by setting negative control values to 0%. Biomix details in Extended Experimental Procedures.

miRNA transfections in immortalized human myoblasts

Immortalized human myoblasts were seeded on 0.4% gelatin in 6-well plates (2.5×105 cells/well) with skeletal muscle proliferation media. Cells were co-transfected with 50nM of indicated miRNA mimics using lipofectamine 2000 (Refer to Table S5). Cells were differentiated with 2% horse serum for 5 days, then lysed as described (Yokota et al., 2009). miRNA delivery was verified via Cy3-labeled CTRL (Life Technologies, AM17120).

Immunofluorescence

7 μm sections of were cut from mouse TAs. Sections were air-dried, hydrated in PBS, and stained for dystrophin as reported (Lu et al., 2000). Image J software (National Institutes of Health), was used for analysis; average pixel intensity was measured after images were set to a 70 pixel threshold and converted to a binary image. Full details in Extended Experimental Procedures.

TNFα treatment of mdx H2K myotubes

H2K myoblasts were differentiated into myotubes in 12-well plates (1.25×105 cells/well) with Matrigel at 37°C. After 4 days of differentiation, myotubes were treated with 1μM VBP15 or prednisolone for 6h, then induced with TNFα (10 ng/mL) for 243h.

Dystrophin quantification with mass spectrometry

Dystrophin protein was quantified using 50 μg of total protein mixed with 25 μg of SILAM internal standard as reported (Brown et al., 2013). Full details in Extended Experimental Procedures.

Animal Studies

All animal studies were done in adherence to the NIH Guide for the Care and Use of Laboratory Animals and experiments were conducted within IACUC guidelines under approved protocols. All studies were reviewed and approved by the Institutional Animal Care and Use Committee of Children’s National Medical Center. Mice were obtained from The Jackson Labs.

Single Myofiber Isolation

Single myofibers were prepared from the EDL muscle of 8-week old mdx and wild-type mice as described (Rosenblatt et al., 1995).

Intramuscular miRNA mimic injections

Six C57BL/10ScSnJ (wild-type) mice aged 6 weeks were injected with 1.5 μg miRNA “Biomix” (Life Technologies, Table S5) or control (Cy3-labeled CTRL miRNA; Life Technologies, AM17120) with tattoo dye to demarcate injection site. Injection was followed by electroporation (2 × 80 V pulses over 20 ms with 980 ms in between pulses) to increase delivery. TAs were harvested after 7 days, snap frozen in liquid-nitrogen cooled isopentane and stored at −80°C.

Notexin-induced muscle damage followed by miRNA mimic injections

Muscle injury was induced by injecting 6 week-old C57BL/10ScSnJ (wild-type) mice with 10 μl of 10 μg/ml notexin (n=3). The TA was surgically exposed (incisions <1cm in length) and tattoo dye marked injection location. Skin was closed with sutures to minimize pain and tissue damage for second injection. Three days later, 10 μg of miRNAs were injected into the right TA; a scrambled Cy3-labeled control mimic (CTRL) was injected into the (left) TA. The muscles were harvested 7 days post-injury.

Systemic delivery of PMOs

Four-week-old mdx mice (C57BL/10ScSn-Dmd<mdx>/J) were given a single 800 mg/kg dose of PMO (Gene Tools): 5′-GGCCAAACCTCGGCTTACCTGAAAT-3′, administered through retro-orbital injection (n=3). Four weeks post-injection (at 8 weeks of age), mice were euthanized via carbon dioxide inhalation; muscles were harvested as described above. Dystrophin protein and miRNA levels were compared to age-matched wild-type controls.

VBP15 and prednisone administration

Archival samples from 2 separate studies were obtained. The first set was from a prophylactic trial where 2-week old mdx mice (C57BL/10ScSn-Dmd<mdx>/J) were dosed with 5 mg/kg prednisolone or 15 mg/kg VBP15 as reported (Heier et al., 2013). The second set was from an “adult trial” where 6-week old mice were dosed with 5 mg/kg prednisolone (53mg/kg) or 45 mg/kg VBP15 for 4 months (Heier et al., 2013). Total RNA was extracted from gastrocnemius (mice sacrificed at 8 weeks) or diaphragm (mice sacrificed at 6 months) and miRNA levels were assessed.

Statistical Analysis

For assays with >2 groups, measurements were compared between groups using one-way ANOVA unless otherwise indicated. Post-hoc linear tests between each group were performed; the resulting p-value reported in figures was adjusted for multiple testing by Sidak method unless otherwise indicated. The contrasting groups in all post-hoc comparisons are indicated in each figure. For assays with two groups where the null hypothesis was testing a change in one direction, a one-tailed, Student’s t-test was utilized, whereas assays where the potential change between groups was + or −, a two-tailed Student’s t-test was utilized. For all bar graphs, data are presented as ± SEM. Details of both tests are specified in the figure legends.

Supplementary Material

1
2

Highlights.

  • miRNAs in muscle microenvironments cause variable dystrophin in muscular dystrophy

  • miRNAs are elevated in dystrophic myofibers and increase with disease severity

  • Inflammatory cytokines induce miRNAs and anti-inflammatories block their expression

  • miRNAs provide a precision medicine target in dystrophy and exon skipping

Acknowledgments

We thank Dr. Vincent Mouly, PhD (Institut de Myologie, Paris, France) for the generous donation of immortalized human myoblasts, and the Eurobiobank, Telethon Network Genetic Biobanks (GTB 12001D) for muscle biopsies. This research was supported by the National Institutes of Health (NIAMS 1P50AR060836-01 Center of Research Translation; NICHD 1U54HD071601-01 Research in Pediatric Developmental Pharmacology Center; and NICHD 1P50AR060836-01 National Center for Medical Rehabilitation Research). Additional funding was provided by an Exploratory Award from Parent Project Muscular Dystrophy. Drs. Fiorillo and Heier were previously supported and Dr. Novak is currently supported by NIAMS training grant 5T32AR056993. Dr. Heier is currently supported by a NHLBI K99 Transition Award K99 HL130035-01A1.

Footnotes

Author Contributions

AAF, EPH, KN, TAP designed and supervised the study. AAF, CRH, CBT, JSN, MCV, KRB, KU, PPN, LB, TAP performed experimental work. JNK provided GRMD and wild-type dog biopsies and characterization. AAF and CRH performed statistical analysis. CA provided the BMD muscle biopsies with deletion characterization. AAF, CRH, KRB performed primary data analysis. AAF, EPH, wrote the manuscript with significant input from CRH, JSN, KN and TAP.

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Supplementary Materials

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NIHMS713383-supplement-1.pdf (1.1MB, pdf)
2
NIHMS713383-supplement-2.docx (16.2KB, docx)

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