RNA Interference Inhibits DUX4-induced Muscle Toxicity In Vivo: Implications for a Targeted FSHD Therapy

Abstract

No treatment exists for facioscapulohumeral muscular dystrophy (FSHD), one of the most common inherited muscle diseases. Although FSHD can be debilitating, little effort has been made to develop targeted therapies. This lack of focus on targeted FSHD therapy perpetuated because the genes and pathways involved in the disorder were not understood. Now, more than 2 decades after efforts to decipher the root cause of FSHD began, this barrier to translation is finally lowering. Specifically, several recent studies support an FSHD pathogenesis model involving overexpression of the myopathic DUX4 gene. DUX4 inhibition has therefore emerged as a promising therapeutic strategy for FSHD. In this study, we tested a preclinical RNA interference (RNAi)-based DUX4 gene silencing approach as a prospective treatment for FSHD. We found that adeno-associated viral (AAV) vector-delivered therapeutic microRNAs corrected DUX4-associated myopathy in mouse muscle. These results provide proof-of-principle for RNAi therapy of FSHD through DUX4 inhibition.

Introduction

Facioscapulohumeral muscular dystrophy (FSHD) is often cited as the third most common muscular dystrophy, affecting 1 in 20,000 individuals.¹ However, recent data indicate FSHD is the most prevalent muscular dystrophy in Europe (1 in 7,500), suggesting its worldwide incidence may be underestimated.² FSHD is an autosomal dominant disorder that typically arises in young adulthood, with most patients showing clinical features before age 30.¹ Age-of-onset and clinical severity vary, but typically all FSHD patients develop progressive and asymmetrical wasting of facial and shoulder girdle muscles. In addition, ~50% of individuals with FSHD display lower limb weakness and ~20% have abdominal pelvic and girdle muscle involvement, which almost invariably leads to hyperlordosis and wheelchair dependence.¹

There is currently no treatment for FSHD, and despite its relative abundance among the muscular dystrophies, very few FSHD-targeted translational studies have been published.³ This historical lack of focus on therapy development was not borne of neglect, but instead arose because the disease is complex and deciphering its pathogenic mechanisms required nearly 2 decades of study.^{4,5,6,7,8,9,10,11,12,13,14,15,16} Although the story is likely not complete, the mechanisms underlying FSHD have started clarifying in recent years, and a new pathogenesis model has emerged.^7,9 Unlike most typical Mendelian disorders, FSHD is not caused by mutations in the coding region of a single gene. Instead, FSHD requires complex genetic and epigenetic changes that conspire to permit expression of one or more myopathic genes.^6,9,10,12,14 Several FSHD candidate genes have been identified, but numerous recent studies support that the primary contributor to FSHD pathogenesis is the pro-apoptotic DUX4 gene, which encodes a transcription factor.^{5,7,8,9,11,13,17} Thus, in the simplest terms, DUX4-overexpression is a primary pathogenic insult underlying FSHD.

The emergence of DUX4 as an important prospective therapeutic target has therefore lowered the barrier to pursuing translational research for FSHD. We hypothesized that reducing DUX4 expression using RNA interference (RNAi) would offer a potential treatment for the disease. In this study, we provide evidence that DUX4 gene silencing, triggered by engineered artificial microRNAs, is myoprotective in vivo. Our data demonstrate proof-of-principle for DUX4 gene silencing as a promising therapeutic approach for FSHD.

Results

Cellular RNAi pathways can be co-opted to suppress dominant disease genes for therapeutic purposes.^{18,19,20,21,22} The goal of this study was to develop a prospective treatment for FSHD using a DUX4-targeted, RNAi-based gene silencing approach. To do this, we engineered five different, mir-30-based artificial microRNAs (miDUX4s; Supplementary Figure S1) targeting the human DUX4 open reading frame. We cloned each miDUX4 sequence into an U6-promoter-driven expression cassette, and performed two in vitro screening assays to identify our lead DUX4-targeted microRNA (Figure 1).^21,23 First, we cotransfected each U6.miDUX4 construct, or a control miRNA vector, with a dual luciferase plasmid in which DUX4 was cloned as the 3′ UTR of Renilla luciferase. A separate Firefly luciferase cassette was used as an internal transfection control (Figure 1a). We then determined the degree of DUX4 gene silencing by measuring the ratio of Renilla to Firefly luciferase activity 24 hours later (Figure 1a). Two constructs (miDUX4.405 and miDUX4.1156) consistently reduced Renilla.DUX4 activity (Figure 1b). We next confirmed their ability to silence DUX4 at the protein level (Figure 1c), and found that miDUX4.405 was consistently superior in vitro. We therefore selected miDUX4.405 as our lead sequence for subsequent in vivo experiments (and heretofore refer to the 405 sequence as miDUX4).

Figure 1.

In vitro screen of miDUX4 sequences. (a) Plasmids used to test DUX4 gene silencing in HEK293 cells. For luciferase assay, the DUX4 cDNA sequences were cloned downstream of the Renilla luciferase (Ren Luc) stop codon, thereby functioning as a 3′ UTR. The Ren Luc.DUX4 fusion transcript was driven by the SV40 promoter. A separate Firefly luciferase cassette was transcribed by the herpes simplex virus thymidine kinase (TK) promoter. CMV indicates cytomegalovirus promoter. (b) Luciferase assay screen to identify lead miDUX4 constructs. DUX4 gene silencing was determined by measuring the ratio of Renilla to Firefly luciferase activity from cells cotransfected with the reporter plasmid and U6 promoter (U6 pro)-driven miDUX4 sequences. (c) Western blot to confirm gene silencing efficacy of miDUX4.405 and miDUX4.1156 following transfection of indicated U6.miRNAs and CMV.DUX4 in HEK293 cells. GAPDH serves as a loading control. UNT indicates untransfected HEK293 cell extracts. GAPDH, glyceraldehyde 3-phosphate dehydrogenase; UTR, untranslated region.

For in vivo gene delivery, we cloned U6.miDUX4 into an adeno-associated viral (AAV) vector separately expressing a CMV.eGFP reporter cassette, and generated AAV6 particles (Figure 2a). No stably expressing DUX4 animal model exists. As a result, we relied upon our previously published AAV-based DUX4 mouse model to express myotoxic levels of DUX4 in wild-type mouse muscles.¹³ Importantly, this model rapidly develops dose-dependent myopathic phenotypes consistent with those reported in other models of muscular dystrophy, including myofiber degeneration and regeneration, fibrosis, muscle weakness, and elevation of cell death pathways.¹³

Figure 2.

DUX4-targeted artificial miRNAs improve DUX4-associated myopathy in vivo. (a) Adeno-associated viral (AAV) vectors used for in vivo studies. Black boxes indicate AAV inverted terminal repeats (ITRs) and pA indicates an SV40 polyadenylation signal. (b) By 2 weeks, control-treated, DUX4-expressing muscles (left) show histopathological evidence of degeneration including: myofibers with central nuclei, inflammatory invasion, and fibrosis. miDUX4-treated muscles (right) are normal at all timepoints. Bar = 50 µm. (c–d) Histological changes were quantified at 3- and 4-weeks postinjection. (c) Shows the percentage of myofibers containing centrally located nuclei (C.N.), an indication of muscle regeneration following damage, 3- and 4-weeks postinjection. (d) Myofiber size distributions in indicated groups. A shift toward small-bore myofibers is characteristic of myopathy. CMV, cytomegalovirus; eGFP, enhanced green fluorescent protein.

To assess the effects of miDUX4 on DUX4-induced myopathy in vivo, we directly injected wild-type C57BL/6 tibialis anterior muscles with an AAV cocktail containing AAV6.DUX4 with either AAV6.miDUX4 or the control vector (CMV.eGFP) (Figure 2a). Control-treated DUX4-expressing muscles showed histological evidence of muscle degeneration and regeneration, 2, 3, and 4 weeks after injection (Figure 2b). In contrast, AAV.miDUX4 protected coinjected DUX4-expressing muscles from degeneration (Figure 2b). Muscles from this group of animals were histologically normal at all timepoints examined (Figure 2b).In addition to obvious visual differences in miDUX4- and control-treated muscles, significant quantifiable changes were evident (Figure 2c,d). Specifically, muscles injected with AAV6.DUX4 alone or AAV6.DUX4 and AAV6.eGFP vectors contained abundant myofibers with reduced size and centrally located nuclei, which are both indications of muscle degeneration and regeneration typically associated with myopathy. In contrast, myofiber sizes and centrally located nuclei were normal in muscles coinjected with AAV6.DUX4 and AAV6.miDUX4 vectors (Figure 2b–d). Improved muscle histology in miDUX4-treated muscles correlated with 90 and 64% average reduction in DUX4 protein and messenger RNA, respectively, compared to uninhibited DUX4 expression in controls (Figure 3a,b).

Figure 3.

DUX4 knockdown in vivo. (a) Western blots demonstrate DUX4 protein knockdown in miDUX4-treated muscles compared to control-treated muscles. DUX4 silencing was evident at all timepoints examined, but only the 2- and 3-week samples are shown here. GAPDH served as a loading control. (b) Real-time PCR shows miDUX4 reduced DUX4 mRNA by 64% in vivo (n = 7 muscles per group from combined 2- and 3-week groups). *Indicates significant difference from AAV.eGFP injected group, P < 0.05 (ANOVA). ANOVA, analysis of variance; eGFP, enhanced green fluorescent protein; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; qPCR, quantitative PCR.

We next determined if AAV.miDUX4 protected muscles from pathological molecular changes associated with FSHD that were downstream of DUX4. For this, we focused on caspase-3, which is expressed in myofibers of FSHD patients and activated by AAV6.DUX4 expression in mouse muscle.^13,24,25 Consistent with our previous results, we found that uninhibited DUX4 expression was associated with caspase-3 positive lesions in AAV6.DUX4-transduced control muscles (Figure 4a). In contrast, there were no caspase-3 positive myofibers in muscles coinjected with AAV6.DUX4 and AAV.miDUX4 vectors (Figure 4b), and caspase-3 transcripts were reduced by 77% in AAV.miDUX4-treated animals (Figure 4c).

Figure 4.

miDUX4-treated myofibers are caspase-3 negative. (a) Serial sections of eGFP control-treated muscle mimics immunohistochemical staining patterns of DUX4-induced myotoxicity indicated by various symbols. Asterisks represent healthy fibers that have DUX4 (+) nuclei but are cleaved caspase-3 (−)/GFP(+). Arrows and arrow heads point to degenerating myofibers: arrows DUX4 (+)/cleaved caspase-3 (+)/GFP(−) and arrow heads DUX4 (+)/cleaved caspase-3 (+)/GFP (+). (b) Treatment with miDUX4 protects myofibers from degeneration indicated by the absence of cleaved caspase-3 (+) myofibers and very few myofibers with DUX4 (+) cytoplasmic staining. Bar = 50 µm. (c) Real-time PCR shows reduced caspase-3 transcript in miDUX4-treated muscles. eGFP, enhanced green fluorescent protein; H&E, hematoxylin and eosin; qPCR, quantitative PCR.

Finally, we measured the effects of AAV6.miDUX4 on DUX4-associated hindlimb grip strength deficits in mice. To do this, we measured forelimb (uninjected) and hindlimb (injected) bilateral grip strengths weekly in wild-type mice following injection with saline or the indicated vectors (Figure 5). By 2 weeks, mice injected with AAV6.DUX4 alone or AAV6.DUX4 with control AAV.eGFP showed significantly reduced grip strength compared to all other groups. This timepoint is consistent with the onset of degeneration in muscle cryosections (Figures 2 and 5). Weakness resolved by 3 weeks, as regenerative processes were underway (Figures 2 and 5). In contrast, animals coinjected with AAV6.DUX4 and AAV6.miDUX4 were not significantly weaker than saline-injected wild-type mice at any timepoint following injection. Mice that received AAV6.miDUX4 alone were unaffected, suggesting that miDUX4 expression was well-tolerated by normal muscles in the absence of a gene target. Together, these results provide evidence for DUX4 inhibition as a therapeutic strategy for FSHD.

Figure 5.

AAV.miDUX4 protects mice from DUX4-associated grip strength deficits. Mice injected with DUX4 and DUX4 with control eGFP show significant hindlimb strength deficits compared to all controls at 2-weeks postinjection. Mice that received DUX4 with miDUX4 treatment were showed no significant difference in strength from controls. Force represents the grip strength ratio of hindlimb (injected) to forelimb (uninjected). N = 8 mice per group; ***P < 0.001 two-way analysis of variance with Bonferroni post-test. eGFP, enhanced green fluorescent protein.

Discussion

Developing treatments for genetic diseases has long been a major goal of biomedical research. Over the years, numerous lethal or debilitating disorders have been transformed into manageable ailments. To date, the list of inherited diseases for which effective treatments exist does not include the muscular dystrophies. Still, much progress has been made in the muscular dystrophy field in the last 3 decades, beginning with the landmark discovery that Duchenne muscular dystrophy (DMD) gene mutations were responsible for DMD in 1986.²⁶ Today, because of similar gene mapping efforts, mutations in at least 40 known genes have been linked to various types of muscular dystrophy. Typically, these disorders arise from mutations that alter the expression, structure, and/or function of a single protein. Identifying a monogenic, protein-coding mutation is an optimal outcome for a gene mapping study, because a relatively straightforward path can be followed toward cell and animal model development, which then provides a platform for testing hypotheses about disease pathogenesis and ultimately developing therapies. For example, DMD gene discovery led to the 1988–1989 identification of the mdx mouse as a DMD model,^27,28 which has been used extensively to study DMD pathogenesis and test therapeutic strategies, some of which are now in the clinical pipeline as prospective DMD treatments.²⁹ This progress would have been impossible without knowledge of the DMD gene mutation or the mdx mouse model.

In contrast to DMD, FSHD translational research has been hindered for decades by uncertainty about disease pathogenesis and the absence of an animal model. As a result, the comparatively few FSHD therapy studies published to date have primarily focused on palliative care, surgical interventions, or treatments that addressed some nonspecific myopathic features common to many types of muscular dystrophy.³ However, the emergence of recent data supporting DUX4 derepression as a key event in FSHD pathogenesis suggests the field has passed a threshold of understanding, making FSHD-targeted translational research more feasible. Despite the recent turn of events, barriers to translation remain. Arguably, the most important such barrier is that a “traditional” FSHD animal model (e.g., one expressing a heritable DUX4 transgene) is still lacking. Following a conservative strategy toward therapeutic development would necessitate delaying in vivo testing of DUX4-targeted approaches until such a model was available. However, this delay would only add to an already potentially long translational path. Indeed, the mdx mouse has been available for nearly 25 years and a DMD “cure” is yet unrealized, so even if a transgenic or knock-in DUX4-expressing FSHD model were in hand today, rapid development of a FSHD therapy would not be guaranteed. Thus, to avoid further delay, we began testing our DUX4 gene silencing approach in vivo using an alternative, AAV-based model of DUX4-associated myopathy, which we previously published.¹³

In our AAV model, DUX4 is expressed at very high levels (using a CMV promoter and myotropic AAV6 capsid) in essentially every myonucleus within a transduced region. The expression levels produced are likely much higher than DUX4 levels detected in biopsies from FSHD patients.^5,11 Indeed, DUX4 so far appears relatively rare in human muscle. One study estimated its presence in only 1 in 1,000–10,000 FSHD myonuclei, and up to 70 cycles of PCR were required to detect the endogenous transcript.¹¹ This apparent scarcity could argue against the involvement of DUX4 in FSHD pathogenesis. Specifically, how could such a rare transcript/protein cause the devastating clinical phenotype sometimes seen in some FSHD patients? This question still remains unresolved in the field. However, one plausible explanation is that a tolerable steady state of DUX4 is permissible in human muscle, and a stochastic series of FSHD-associated DUX4 amplifications produce small, myopathic events that cumulatively lead to significant muscle loss over time. Thus, observing a rare DUX4 amplification event in a small muscle biopsy sample would be difficult at any single timepoint.

The dose dependency of DUX4 toxicity lends support to this amplification model. Indeed, in a previous study we found that high doses of AAV6.DUX4 produced widespread lesions throughout an entire mouse tibialis anterior muscle, but damage manifested more slowly and focally at decreasing dosages.¹³ In this study, DUX4 gene silencing by AAV.miDUX4 protected muscles from DUX4-induced damage, even though the protein was still detectable in virtually all transduced myonuclei (Figure 4). In addition, our in vivo work here is consistent with a recently published study showing DUX4-targeted siRNA or antisense RNA could inhibit an atrophic phenotype in cell culture.³⁰ Together, these studies support the feasibility of DUX4 inhibition by RNAi as a potential therapy for FSHD.

Before moving these preclinical studies forward, issues common to other nucleic acid-based therapies must be addressed. One is safety. RNAi therapy is still a new and clinically unproven approach. At very high doses, exogenous inhibitory RNAs may sometimes saturate natural miRNA biogenesis pathways, thereby potentially causing nonspecific cytotoxicity. Sequence-specific off-target effects can also occur if an inhibitory RNA shares complementary sequence with other, unintended transcripts.^19,31 However, overexpression related toxicity was associated with suboptimally processed, first generation short hairpin RNA (shRNA) vectors, while the miRNA-based approach we used here generates a transcript that cycles through the miRNA biogenesis pathway efficiently and thus avoids this toxicity.¹⁹ Importantly, our proof-of-principle studies here already support the safety of our approach, as high doses of AAV6.miDUX4 vector produced no overt functional pathology in wild-type mice lacking a DUX4 target (Figure 5). Moreover, bioinformatics analyses support that our miDUX4 sequences are DUX4-specific and should not directly silence off-target transcripts. Additional dose escalation studies and off-targeting analyses in mice, human cells, and perhaps non-human primates, will further solidify the safety of our miDUX4 vectors.

A second major issue that our strategy shares with other nucleic acid therapies is delivery. Currently, AAV vectors are the tool of choice for muscle gene delivery because of their myotropism and excellent safety profile. Indeed, AAV has already proven safe and effective in human clinical trials,^32,33 supporting that our approach potentially has immediate practicability if myofibers are the only DUX4-expressing target cells. However, it is difficult to predict the effectiveness of our AAV.miDUX4 approach should muscle progenitor cells also require DUX4 inhibition. This uncertainty arises from the lack of data about AAV tropism in muscle satellite cells (SC). It will therefore be important to determine if FSHD pathology is related to DUX4 expression in SCs, and if so, whether SC transduction is possible with AAV6 or another AAV serotype. If DUX4 suppression in SCs is required, and AAV vectors are non-transducing, it may be possible to rationally design or pan for SC-tropic AAV capsids to enhance efficacy.^34,35 Alternatively, in vitro-synthesized inhibitory RNAs could be used as cell-penetrating delivery strategies continue to improve.

A final potential complicating issue related to a DUX4-targeted therapy is the necessity for allele-specific silencing. Specifically, DUX4 can be alternatively spliced to produce a short, apparently non-toxic form lacking the C-terminal transactivation domain (DUX4-s).¹¹ Although the function of DUX4-s is uncertain, and it too seems to be expressed at similarly low levels (like full-length DUX4; DUX4-fl) in muscle, there is a hypothesis currently in the field that DUX4-s contributes some important function to normal muscle, while preferential expression of the full-length DUX4 isoform in FSHD contributes to myopathic phenotypes. If true, it may be necessary to specifically silence only DUX4-fl. Importantly, some of our miDUX4 constructs can accomplish this allele-specificity, including miDUX4.1156 (Figure 1b,c).

In conclusion, we have entered an exciting period in FSHD research. New basic discoveries on FSHD pathogenesis have opened the door to begin developing, arguably for the first time, FSHD-specific therapeutic strategies. Such approaches will likely include DUX4 inhibition, as we described here using RNAi. Nevertheless, although the pathogenic mechanisms underlying FSHD are clarifying, the picture is likely still incomplete. Our current and previous studies²¹ set the stage for applying RNAi more broadly to other FSHD-associated gene targets, as they emerge.

Materials and Methods

Cloning of DUX4-targeted microRNAs. Five different mouse U6 promoter-driven artificial microRNAs targeting human DUX4 (called miDUX4s) were cloned as previously described.^21,23 Each U6.miDUX4 construct was derived from human mir-30 stem and loop sequences and structures, but the 22 nucleotide (nt) mature mir-30 duplex was replaced by sequences targeting the DUX4 gene (Supplementary Figure S1). The miDUX4 nomenclature refers to the first position of the miRNA binding site relative to the +1 of the DUX4 coding region (MAL start). The control U6-driven artificial miRNA targeting eGFP (miGFP) was previously described.²¹

Luciferase assay. The luciferase reporter plasmid (Figure 1a) was modified from Psicheck2 (Promega, Madison, WI).²³ Human DUX4 cDNA was cloned downstream of the Renilla luciferase stop codon, thereby functioning as a 3′ UTR. A separate thymidine kinase (TK) promoter-driven Firefly luciferase cassette, present on the same plasmid, served as a transfection control. HEK293 cells were cotransfected in triplicate wells (Lipofectamine-2000; Invitrogen, Carlsbad, CA) with the luciferase DUX4 reporter and individual U6.microRNA expression plasmids in a 1:5 molar ratio. DUX4 gene silencing was determined by measuring Renilla and Firefly luciferase activity (Dual Luciferase Reporter Assay System; Promega) 24 hours post-transfection, following manufacturer's instructions. Triplicate data were averaged, and individual experiments performed three times; results were reported as the mean ratio of Renilla to Firefly activity ± SEM.

Western blot. For in vitro work, HEK293 cells were cotransfected with U6.miDUX4 or control microRNA plasmids and a CMV.DUX4 expression vector at an 8:1 molar ratio. Protein was extracted 48 hours later (M-PER from Pierce, Rockford, IL). For in vivo work, protein was extracted from muscles injected 1.5–4 weeks prior, using previously described methods. Protein was quantified by Lowry assay (BioRad, Hercules, CA), 30 µg samples were separated on 12% SDS-PAGE, transferred to polyvinylidene fluoride membrane, and incubated with the following antibodies: mouse monoclonal antibody to V5 (HRP-coupled) (1:5,000; Invitrogen); mouse monoclonal GAPDH antibody (1:500; Millipore, Billerica, MA) overnight at 4 °C. GAPDH-probed blots were washed, then incubated with HRP-coupled goat anti-mouse secondary antibody (1:100,000; Jackson ImmunoResearch, West Grove, PA) for 1 hour at room temperature. Following washes, blots were developed using Immobilon Western HRP substrate (Millipore), and exposed to film.

AAV vector delivery to mouse muscle. U6.miDUX4.405 was cloned into AAV.CMV.eGFP (aka AAV.eGFP) proviral plasmid upstream of CMV.eGFP. AAV6.DUX4 (aka AAV.CMV.DUX4) was previously described.¹³ AAV6 particles were generated and titrated as previously described by the Viral Vector Core Facility at The Research Institute at Nationwide Children's Hospital. Eight-week-old C57BL/6 female mice received 50 µl direct intramuscular injections into the tibialis anterior. For all experiments except grip strength, premixed virus cocktails contained 8 × 10⁸ DNAse resistant particles of AAV6.DUX4 and 3 × 10¹⁰ of either AAV6.miDUX4 or AAV6.eGFP. For grip strength studies, animals received 3 × 10⁹ DNAse resistant particles of AAV6.DUX4. All mouse procedures were performed following guidelines approved by the Institutional Animal Care and Use Committee (IACUC) at the Research Institute at Nationwide Children's Hospital.

Imaging and histology. In vivo AAV transduction was determined by eGFP epifluorescence using a fluorescent dissecting microscope (MZ16FA; Leica, Wetzlar, Germany) at ×4.63 magnification. Dissected muscles were placed in O.C.T. Compound (Tissue-Tek, Torrance, CA), frozen in on liquid nitrogen-cooled isopentane, cut onto slides as 10 µm cryosections, and stained with hematoxylin and eosin (following standard protocols),¹³ V5 (Millipore) and cleaved Caspase-3 (Cell Signaling Technology, Danvers, MA) polyclonal antibodies. For immunohistochemistry, cryosections were postfixed in 4% paraformaldehyde, washed, blocked in phosphate-buffered saline with 5% goat serum and 0.3% Triton X-100, incubated overnight at 4 °C with primary antibody (cleaved Caspase-3 1:1,500 and V5 1:1,000; in 1% bovine serum albumin, 0.3% Triton X-100, and phosphate-buffered saline), and then with AlexaFluor-594 conjugated goat anti-rabbit secondary antibodies (1:500; 1 hour at room temperature; Molecular Probes, Carlsbad, CA). Slides were covered in Vectashield plus DAPI (Vector Labs, Burlingame, CA). Muscle cross-sectional fiber diameters and percentage of myofibers with centrally located nuclei were determined from muscles injected 3 and 4 weeks prior (n = 4 muscles per group; five representative ×20 photomicrographs per section), using cellSens Version 1.3 software (Olympus, Center Valley, PA).

Real-time PCR. RNA was extracted from muscles 2–3 weeks postinjection(N = 7 muscles per treatment, TRI Reagent; Molecular Research Center, Cincinnati, OH; followed by RNeasy mini column purification; Qiagen, Valencia, CA). RNA was quantified (NanoDrop ND-1000 Spectrophotometer; Thermo Scientific, Wilmington, DE) and DNAse treated for 30 minutes at 37 °C (DNA-free; Ambion, Foster City, CA). Following DNAse inactivation, RNA was reverse transcribed using random hexamers (High Capacity cDNA Reverse Transcription Kit; Applied Biosystems, Foster City, CA) and gene specific primers to the DUX4 coding region (5′-GCTAGCCACCATGGCCCTCCCGACAC-3′) and the DUX4 coding region with an additional tag sequence (5′-CGACTGGAGCACGAGGACACTGACGATGCCCGGGTACGGGTTCCGCTCAAAGC-3′).¹¹ Quantitative real-time PCR was performed using a SYBR green reaction (SA Biosciences, Frederick, MD)for DUX4 (5′-GCTAGCCACCATGGCCCTCCCGACAC-3′ and 5′-CGACTGGAGCACGAGGACACTGA-3′)and GAPDH (5′-CACGGCAAATTCAACGGCACAGTCAAGG-3′ and 5′-GTTCACACCCATCACAAACATGG-3′).¹¹ Caspase-3 levels were detected using a TaqMan Assay primer/probe set (Applied Biosystems) and normalized to GAPDH. All samples were run in triplicate.

Grip strength. Grip strength was measured in forelimbs and hindlimbs of C57BL/6 mice 1 week before injection to establish a baseline, and then weekly up to 4-weeks postinjection as previously described (n = 7 animals per group).¹³ Data represent mean of hindlimb force divided by forelimb force ± SEM.

SUPPLEMENTARY MATERIAL Figure S1. miDUX4 structures, sequence, and binding sites.

Supplementary Material

Figure S1.

miDUX4 structures, sequence, and binding sites.