Mutations in the dystrophin gene cause Duchenne muscular dystrophy (DMD), which is characterized by lethal degeneration of cardiac and skeletal muscles. Mutations that delete exon 44 of the dystrophin gene represent one of the most common causes of DMD and can be corrected in ~12% of patients by editing surrounding exons, which restores the dystrophin open reading frame. Here, we present a simple and efficient strategy for correction of exon 44 deletion mutations by CRISPR-Cas9 gene editing in cardiomyocytes obtained from patient-derived induced pluripotent stem cells and in a new mouse model harboring the same deletion mutation. Using AAV9 encoding Cas9 and single guide RNAs, we also demonstrate the importance of the dosages of these gene editing components for optimal gene correction in vivo. Our findings represent a significant step toward possible clinical application of gene editing for correction of DMD.

The second most common mutational hotspot in the dystrophin gene includes exon 44, which disrupts the open reading frame in surrounding exons ( 4 , 7 – 9 ). Here, we describe the creation of a new mouse model of DMD with exon 44 deletion, and we present two strategies for correction of this mutation by CRISPR-Cas9–mediated skipping of surrounding exons. These mice represent an important tool for the testing and optimization of diverse therapies for DMD. We also show that sgRNAs, unexpectedly, are limiting for optimal gene editing in vivo and that editing efficiency can be enhanced ~10-fold by optimizing the dose of AAVs encoding Cas9 and sgRNAs. Our findings highlight the potential of gene editing to permanently eradicate mutations that cause DMD, thereby preventing the pathogenic sequelae of this disease.

A substantial challenge in the development of DMD therapies has been the lack of animal models harboring the most common human mutations. Because the mouse and human dystrophin genes both contain 79 exons with highly conserved exon splicing patterns, results obtained in mouse models of the disease can be extrapolated to the human condition. One of the most common deletions in patients with DMD eliminates exon 50 in the rod domain of dystrophin, which places exon 51 out of frame with preceding exons ( 4 , 7 – 10 ). We recently described the rescue of mice and dogs lacking exon 50 by injection of two adeno-associated viruses of serotype 9 (AAV9) encoding the CRISPR-Cas9 gene and single guide RNAs (sgRNAs) that allow skipping or reframing of exon 51 and restoration of dystrophin expression ( 11 , 12 ).

Duchenne muscular dystrophy (DMD), caused by mutations in the dystrophin gene, is characterized by degeneration of cardiac and skeletal muscles, loss of ambulation, and premature death ( 1 ). Dystrophin is a massive protein (>3600 amino acids), which stabilizes muscle membranes by tethering the actin cytoskeleton to the inner surface of the sarcolemma ( 2 , 3 ). Thousands of mutations that prevent dystrophin production have been identified in patients with DMD ( 4 ). These mutations cluster in hotspot regions of the gene that can, in principle, be bypassed by various exon skipping strategies to restore the dystrophin open reading frame ( 5 ). To date, however, there has been no effective long-term therapy for this disease, and the only drug approved by the Food and Drug Administration for the treatment of DMD allows for restoration of <1% of the normal level of dystrophin protein after extended treatment ( 6 ). Thus, there remains a major unmet medical need for new strategies to correct the underlying cause of DMD—genetic mutations in the dystrophin gene.

To examine the effect of dystrophin restoration on muscle function in systemically corrected ΔEx44 DMD mice, we performed electrophysiology on the EDL muscle of ΔEx44 DMD mice at 4 weeks after injection with AAV-Cas9 and AAV-G6. We observed rescue of maximal tetanic force in the EDL muscle of the corrected ΔEx44 DMD mice ( Fig. 4C ). Improvement of muscle function correlated with increased dystrophin expression and decreased muscle degeneration and was associated with administration of increasing amounts of AAV-G6 relative to AAV-Cas9 (fig. S9). For measurement of muscle-specific force, which is calibrated with the muscle cross-sectional area, we observed an increase in force from 59 to 89% for a 1:5 ratio and to 107% for a 1:10 ratio of AAV-Cas9:AAV-G6 in the EDL muscle of systemically corrected ΔEx44 DMD mice ( Fig. 4D ). We conclude that systemic delivery of AAV-Cas9 and AAV-G6 efficiently restores dystrophin expression and improves muscle function in corrected ΔEx44 DMD mice, and the amount of sgRNA delivered to the muscle is critical to the efficiency of genome editing in vivo.

To further assess systemic delivery of AAV-Cas9 in the presence of different amounts of AAV-G6, we performed Western blot analysis to evaluate the amount of Cas9 protein expressed in the muscles. Although we kept the total AAV-Cas9 dosage constant (5 × 10 13 vg/kg), the mice that received higher doses of AAV-G6 showed greater expression of Cas9 protein in corrected muscles ( Fig. 4B and fig. S6). Quantitative polymerase chain reaction (qPCR) analysis of Cas9 mRNA in muscle groups comparing low and high doses of AAV-G6 revealed increased Cas9 mRNA expression in the skeletal muscle in the presence of high doses of AAV-G6 (fig. S8, C and D). These results indicate that Cas9 expression is affected by the amount of sgRNA present, and thus, sgRNA is limiting for optimal gene editing in vivo. These results also suggest that the extent of dystrophin restoration and muscle recovery may provide an environment that favors Cas9 expression.

( A ) Immunostaining shows restoration of dystrophin in the TA, triceps, diaphragm, and heart of ΔEx44 mice 4 weeks after systemic delivery of AAV-Cas9 and AAV-G6 at the indicated ratios. Dystrophin is shown in red. Nuclei are marked by DAPI stain in blue. Scale bar, 100 μm ( B ) Western blot analysis shows restoration of dystrophin expression in the TA, triceps, diaphragm, and heart of ΔEx44 mice 4 weeks after systemic delivery of AAV-Cas9 and AAV-G6 at the indicated ratios. Vinculin the loading control (n = 4). ( C ) Maximal tetanic force of the EDL muscles in WT (blue), ΔEx44 DMD (red), and corrected ΔEx44 DMD (green) mice 4 weeks after systemic delivery of AAV-Cas9 and AAV-sgRNA at 1:5 and 1:10 ratios. P < 0.005 (n = 6). ( D ) Specific force (mN/mm 2 ) of the EDL muscles in WT (blue), ΔEx44 DMD (red), and corrected ΔEx44 DMD (green) mice 4 weeks after systemic delivery of AAV-Cas9 and AAV-sgRNA at 1:5 and 1:10 ratios. Data are represented as mean ± SEM. One-way ANOVA was performed, followed by Newman-Keuls post hoc test. **P < 0.001 (n = 6).

To achieve body-wide rescue of the disease phenotype in ΔEx44 DMD mice, we delivered AAV-Cas9 and AAV-G6 systemically through intraperitoneal injection. AAV-Cas9 was injected at a dosage of 5 × 10 13 vg/kg. Multiple ratios of AAV-G6 to AAV-Cas9 were tested to determine whether there might be an optimal ratio of the viruses for maximal systemic editing efficiency. Four weeks after injection, we assessed dystrophin protein expression in several muscle tissues, including TA muscle of the hindlimb, triceps of the forelimb, diaphragm, and cardiac muscle. By immunostaining, we observed dystrophin expression in 94, 90, and 95% of myofibers in the TA, triceps, and diaphragm, respectively, and in 94% of cardiomyocytes when ΔEx44 mice were injected with a 1:10 ratio of AAV-Cas9:AAV-G6 (5 × 10 13 vg/kg of AAV-Cas9 and 5 × 10 14 vg/kg of AAV-G6) ( Fig. 4A and fig. S5). The restoration of dystrophin protein in skeletal muscles correlated with the dosage of AAV-G6 delivered through intraperitoneal injection. In contrast, in the heart, dystrophin-positive cardiomyocytes were seen at a low dosage of AAV-G6 and remained consistent at higher dosages. Western blot analysis of the same muscle groups after systemic delivery showed similar trends of dystrophin correction ( Fig. 4B and fig. S6). At every ratio of AAV-Cas9:AAV-G6 tested by systemic delivery, the cardiac muscle showed higher dystrophin restoration than the skeletal muscle. Correction of the cardiac muscle reached 82% when injected at a 1:1 ratio of AAV-Cas9:AAV-G6 and increased an additional 12% at a 1:10 ratio. In contrast, we observed an increase in dystrophin-expressing myofibers from 10 to 94% when the sgRNA was increased. H&E and Picrosirius red staining showed that histopathologic hallmarks of muscular dystrophy, such as regenerated fibers with central nuclei, were diminished in the TA, diaphragm, and triceps muscles at 4 weeks after AAV-Cas9 and AAV-G6 delivery (figs. S7 and S8A). Quantitative analysis of the distribution of muscle fiber cross-sectional area showed an improvement in the TA muscle at 4 weeks after delivery of AAV-Cas9:AAV-G6 at 1:5 and 1:10 ratios (fig. S8B).

On the basis of the CRISPR design tools ( http://crispr.mit.edu/ and https://benchling.com/ ), we determined the top 10 potential off-target sites, and on the basis of sequencing analysis, we did not detect off-target effects at these sites (fig. S4). The T7E1 analysis confirmed the absence of off-target cutting in the top 10 potential off-target sites, and DNA sequencing of the isolated genomic PCR amplification products spanning the potential off-target sites confirmed the absence of sgRNA/Cas9-mediated mutations at the predicted sites (fig. S4A). In addition, we performed genomic amplicon deep sequencing of the top 10 predicted off-target sites within protein-coding exons. None of these sites showed significant sequence alterations (fig. S4, B and C).

To evaluate dystrophin protein restoration after intramuscular injection with AAV-Cas9 and AAV-G5 or AAV-G6, we performed Western blot analysis on the TA muscle and the heart ( Fig. 3D ). We observed restoration of dystrophin protein expression to 74% of the WT level in edited TA muscles of ΔEx44 DMD mice ( Fig. 3E ). Although the injection was localized to the TA muscle, we observed expression of dystrophin in the heart at 21% of the WT level ( Fig. 3D ). This suggests leakage of AAV into the circulation and delivery of the gene editing components to the heart. Immunostaining showed that dystrophin protein expression was restored in 99% of the myofibers in TA muscle injected with AAV-Cas9 and AAV-G6 ( Fig. 3F and fig. S3F). Histological analysis and hematoxylin and eosin (H&E) staining showed a pronounced reduction in fibrosis, necrotic myofibers, and regenerating fibers with central nuclei, indicating amelioration of the abnormalities associated with muscular dystrophy in the TA muscle 3 weeks after AAV9-Cas9 and AAV-G6 injection ( Fig. 3G ).

Genomic and cDNA amplicon deep sequencing on the target region of the TA muscles with AAV-G6 intramuscular injection also confirmed that 9.8% of mutations at the genomic level and 35.7% of mutations at the mRNA level contain a single A insertion at the cutting site after gene editing with AAV-G6 (fig. S3, D and E). This single A insertion leads to reframing of exon 45 and restores the dystrophin protein reading frame. We also observed minor AAV inverted terminal repeat (ITR) integration events at the cutting site, with a frequency of 0.21% at the genomic level (fig. S3D) and 1.15% at the mRNA level (fig. S3E).

To further evaluate the mutations generated by gene editing, we performed topoisomerase-based thymidine to adenosine (TOPO-TA) cloning using the RT-PCR amplification products and sequenced the cDNA products. Sequencing results demonstrated that 7% of sequenced clones represented exon 45–skipped cDNA products, and 42% of sequenced clones contained a single adenosine (A) insertion in exon 45 that resulted in reframing of dystrophin protein ( Fig. 3, B and C ). The predominance of reframing explains the high abundance of the RT-PCR band at 355 bp and the lower abundance of the smaller RT-PCR product at 179 bp, which reflects exon skipping ( Fig. 3A ).

( A ) RT-PCR analysis of TA muscles from WT and ΔEx44 mice 3 weeks after intramuscular injection of gene editing components carried by AAV9. Lower dystrophin bands (179 bp) indicate skipping of exon 45. ( B ) Pie chart showing percentage of events detected at exon 45 after AAV-Cas9 and AAV-G6 treatment using RT-PCR sequence analysis of TOPO-TA generated clones. RT-PCR products were divided into four groups: NE, not edited; SK, exon 45 skipped; RF, reframed; and OF, out of frame. Data are represented as mean ± SEM (n = 3). ( C ) Sequences of RT-PCR products of WT, ΔEx44, and corrected ΔEx44 mice. In-frame sequences are shown in blue, including WT and exon 45–skipped sequences. Reframed sequence is shown in green, and out-of-frame sequence is shown in red. ( D ) Western blot analysis shows restoration of dystrophin expression in the TA muscle and heart of ΔEx44 mice. Vinculin is the loading control. ( E ) Quantification of the Western blot analysis in the TA muscle. Relative dystrophin intensity was calibrated with vinculin internal control. Data are represented as mean ± SEM. Unpaired Student’s t test was performed. *P < 0.005 (n = 3). ( F ) Immunostaining shows restoration of dystrophin in the TA muscle of ΔEx44 mice 3 weeks after intramuscular injection of gene editing components carried by AAV9. Dystrophin is shown in red. Nuclei are marked by DAPI stain in blue. Scale bar, 100 μm (n = 3). ( G ) H&E staining of the TA and heart in WT, ΔEx44, and corrected ΔEx44 mice. Inset boxes indicate areas of magnification shown below. Scale bars, 50 μm (n = 3).

To validate the efficacy of the single-cut gene editing strategy in the ΔEx44 DMD mouse model, we performed localized intramuscular injection of AAV9 encoding SpCas9 (AAV-Cas9) and AAV9 encoding sgRNA (AAV-G5 or AAV-G6) in the TA muscle of postnatal day 12 (P12) mice. As a control group, WT and ΔEx44 DMD mice were injected with AAV-Cas9 without AAV-sgRNA. In initial studies, 50 μl of AAV9 (1 × 10 12 vg/ml) was injected per leg, containing equal amounts of AAV-Cas9 and AAV-G5 or AAV-G6. Three weeks after the intramuscular injection, we collected the TA muscles for analysis. In vivo gene editing by AAV-G5 and AAV-G6 was compared by the T7E1 assay and RT-PCR of the targeted region ( Fig. 3A and fig. S3C). Gene editing with AAV-G6 showed higher efficiency based on DNA cutting in vivo (fig. S3C). RT-PCR with primers that amplify the region from exon 43 to exon 46 revealed deletion of exon 45 in the TA muscle injected with AAV-Cas9 and AAV-G6 ( Fig. 3A ). This allows exon 43 to skip exon 45 and directly splice to exon 46 when processing the pre-mRNA. As a result, the alternate mRNA enables the production of a truncated dystrophin protein in the corrected TA muscle of ΔEx44 DMD mice.

We first compared the efficiency of gene editing with different expression constructs encoding Cas9 and sgRNAs. The PX458 plasmid encodes both editing components ( 21 ), whereas we used two AAV expression plasmids to express Cas9 and three copies of the sgRNA that targeted exon 45 in mouse C2C12 muscle cells. By using the T7E1 assay, we observed comparable editing efficiency with both constructs (fig. S3B). Among the two sgRNAs tested, G6 showed better cutting efficiency than G5, consistent with the observations in mouse 10T½ cells and human 293 cells (fig. S1C).

To deliver SpCas9 and sgRNA in vivo, we used AAV9 to package the gene editing components. AAV9 is a single-stranded DNA virus that displays tropism to both skeletal muscle and heart and has been used in numerous clinical trials ( 14 – 17 ). To further achieve muscle-specific gene editing, we used the creatine kinase 8 (CK8e) regulatory cassette that combines key elements of the enhancer and promoter regions of the muscle CK gene to drive SpCas9 expression in skeletal muscle and heart ( 18 , 19 ). For the delivery of sgRNA, we used three RNA polymerase III promoters (U6, H1, and 7SK) to express three copies of the sgRNA (fig. S3A) ( 20 ).

Shear force generated during muscle contraction leads to muscle membrane tearing in muscle lacking dystrophin, eventually causing myofiber degeneration and muscle fibrosis ( 13 ). Fibrotic tissue increases muscle stiffness and compromises contractility of muscles. To further analyze muscle function of ΔEx44 DMD mice, we measured maximal tetanic force in the extensor digitorum longus (EDL) muscle ex vivo. Compared with WT littermates at 4 weeks of age, ΔEx44 DMD mice showed an ~50% decrease in the specific and absolute tetanic force in the EDL muscle ( Fig. 2, H and I ). A similar decrease in muscle strength was observed by grip strength analysis in 8-week-old ΔEx44 DMD mice ( Fig. 2J ).

( A ) CRISPR-Cas9 editing strategy used for generation of mice with exon 44 deletion (ΔEx44). Exon 45 (red) is out of frame with exon 43. ( B ) RT-PCR analysis of TA muscles to validate deletion of exon 44. RT-PCR primers were in exons 43 and 46, and the amplicon size is 503 bp for WT mice and 355 bp for ΔEx44 DMD mice. RT-PCR products are schematized on the right (n = 3). ( C ) Sequencing of RT-PCR products from ΔEx44 DMD mouse muscle confirmed deletion of exon 44 and generation of a premature stop codon in exon 45, indicated by red asterisk. ( D ) Dystrophin staining of the TA, diaphragm, and heart of WT and ΔEx44 DMD mice. Dystrophin is shown in red. Nuclei are marked by DAPI stain in blue. Scale bar, 100 μm. ( E ) The Western blot analysis shows loss of dystrophin expression in the TA, gastrocnemius/plantaris (G/P) muscle, and heart of ΔEx44 mice. Vinculin is the loading control (n = 3). ( F ) H&E staining of the TA, diaphragm, and heart. Note extensive inflammatory infiltrate and centralized myonuclei in ΔEx44 sections. Inset boxes indicate areas of magnification shown below. Scale bars, 50 μm. ( G ) Serum creatine kinase (CK), a marker of muscle damage and membrane leakage, was measured in WT (C57BL/6 and C57BL/10), ΔEx44, and mdx mice. Data are represented as mean ± SEM. Unpaired Student’s t test was performed. *P < 0.005 (n = 6). ( H ) Representative trace of maximal tetanic force of EDL muscles in WT (blue) and ΔEx44 mice (red). P < 0.005 (n = 6). ( I ) Specific force of EDL muscles in WT (blue) and ΔEx44 mice (red). Data are represented as mean ± SEM. Unpaired Student’s t test was performed. **P < 0.001 (n = 6). ( J ) Forelimb grip strength analysis of WT and ΔEx44 mice. Data are represented as mean ± SEM. Unpaired Student’s t test was performed. **P < 0.001 (n = 6).

To optimize gene editing for correction of an exon 44 deletion in vivo, we generated a mouse model bearing an exon 44 deletion in the Dmd gene by CRISPR-Cas9 gene editing ( Fig. 2A ). We injected zygotes of C57BL/6 mice with two sgRNAs that target the introns flanking exon 44 and implanted the zygotes into surrogate female mice (fig. S2A). An F0 founder with a 908-bp deletion that eliminated exon 44 was chosen for further studies. These ΔEx44 DMD mice contain one of the most common deletions responsible for DMD in humans. In principle, correction of exon 44 deletions by gene editing of surrounding exons could potentially restore the reading frame of dystrophin in ~12% of patients with DMD. Deletion of exon 44 was confirmed by reverse transcription polymerase chain reaction (RT-PCR) analysis ( Fig. 2B ). Sequencing of the RT-PCR products using primers for sequences in exons 43 and 46 confirmed the removal of exon 44 in these mice ( Fig. 2C ). At 4 weeks of age, immunostaining of the tibialis anterior (TA) muscle, diaphragm, and heart in the ΔEx44 DMD mice showed complete absence of dystrophin protein expression ( Fig. 2D ). Western blot analysis confirmed loss of dystrophin protein ( Fig. 2E ). Fibrosis, inflammatory infiltration, and regenerative fibers with centralized nuclei were observed in 4-week-old ΔEx44 DMD mice, indicative of a severe muscular dystrophy phenotype ( Fig. 2F and fig. S2B). Serum creatine kinase (CK) levels in the ΔEx44 DMD mice were elevated 22-fold compared with wild-type (WT) littermates, similar to mdx mice, an established DMD mouse model ( Fig. 2G ).

Because of the high efficiency of editing in the T7E1 assay and the complete conservation of sequence between human and mouse genomes, we chose to use sgRNA G6 to derive single clones of ΔEx44 iPSCs that were edited within exon 45. Thirty-four single clones were isolated and expanded. Sequence analysis of the clones showed exon skipping events in 3 of 34 clones, and dystrophin reframing by either +3n+1 or +3n−2 in 13 of 34 clones (fig. S1D). Western blot analysis confirmed the restoration of dystrophin expression in three of the CRISPR-Cas9 corrected clones (fig. S1E).

sgRNAs with the highest gene editing activity based on the T7E1 assays were then tested for the ability to efficiently edit the corresponding exons in patient-derived iPSCs lacking exon 44 (referred to as ΔEx44). A single plasmid encoding optimized sgRNAs (G3 or G4 for exon 43, or G6 for exon 45) and Streptococcus pyogenes Cas9 (SpCas9) was introduced into ΔEx44 patient-derived iPSCs by electroporation, and the edited iPSCs were differentiated into cardiomyocytes (iPSC-CMs). Dystrophin expression was assessed by Western blot analysis and immunostaining, confirming restoration of dystrophin protein expression in edited ΔEx44 iPSC-CMs ( Fig. 1, E and F ). Levels of dystrophin protein expression in ΔEx44 iPSC-CMs edited with sgRNAs G4 and G6 were approximately comparable to those seen in healthy control iPSC-CMs ( Fig. 1E ).

We selected sgRNAs that permit deletion of the splice acceptor or donor sites of exons 43 and 45, thereby allowing splicing between surrounding exons to recreate in-frame dystrophin. For editing exon 43, we designed four 20–nucleotide (nt) sgRNAs (G1, G2, G3, and G4) directed against sequences near the 5′ and 3′ boundaries of the splice junctions of exon 43 ( Fig. 1C ). For exon 45, we observed that the intron-exon junction of the splice acceptor site is contained within a 33–base pair (bp) region that is identical in the human and mouse genomes, allowing exon skipping strategies to be interchanged between the two species (fig. S1A). We generated four 18- to 20-nt sgRNAs (G5, G6, G7, and G8) to target the 5′ boundary of exon 45 within the conserved region of the human and mouse genomes ( Fig. 1D ). By the mismatch-specific T7 endonuclease I (T7E1) assay, we compared the sgRNAs for their ability to direct Cas9-mediated gene editing in human 293 cells (fig. S1B). Two of four sgRNAs for exon 43 efficiently edited the targeted region, and all four sgRNAs for exon 45 generated precise cuts at the conserved region (fig. S1C). We concurrently tested the editing activity of the same four sgRNAs for exon 45 in mouse 10T½ cells and confirmed the effectiveness of the four sgRNAs in both the human and mouse genomes (fig. S1C).

( A ) Schematic of the procedure for deriving and editing patient with DMD–derived iPSCs and iPSC-CMs. ( B ) Gene editing strategy for DMD exon 44 deletion. Deletion of exon 44 (black) results in splicing of exons 43 to 45, generating an out-of-frame stop mutation of dystrophin. Disruption of the splice junction of exon 43 or exon 45 results in splicing of exons 42 to 45 or exons 43 to 46, respectively, and restores the protein reading frame. The protein reading frame can also be restored by reframing exon 43 or 45 (green). ( C ) Sequence of sgRNAs targeting exon 43 splice acceptor and donor sites in the human DMD gene. The protospacer adjacent motif (PAM) (denoted as red nucleotides) of the sgRNAs is located near the exon 43 splice junctions. Exon sequence is represented by letters in bold uppercase. Intron sequence is represented by letters in lowercase. Arrowheads show sites of Cas9 DNA cutting with each sgRNA. Splice acceptor and donor sites are shaded in yellow. ( D ) Sequence of sgRNAs targeting exon 45 splice acceptor site in the human DMD gene. The PAM (denoted as red nucleotides) of the sgRNAs is located near the exon 45 splice acceptor site. The human and mouse conserved sequence is shaded in light blue. Exon sequence is represented by letters in bold uppercase. Intron sequence is represented by letters in lowercase. ( E ) Western blot analysis shows restoration of dystrophin expression in exon 43–edited (E43) and exon 45–edited (E45) ΔEx44 patient iPSC-CMs with sgRNAs (G) 3, 4, and 6, as indicated. Vinculin is the loading control. HC indicates iPSC-CMs from a healthy control. The second lane is the unedited ΔEx44 patient iPSC-CMs. ( F ) Immunostaining shows restoration of dystrophin expression in exon 43–edited and exon 45–edited ΔEx44 patient iPSC-CMs. Dystrophin is shown in red. Cardiac troponin I is shown in green. Nuclei are marked by 4′,6-diamidino-2-phenylindole (DAPI) stain in blue. Scale bar, 50 μm.

We generated patient-derived induced pluripotent stem cells (iPSCs) from a patient with DMD lacking exon 44 of the dystrophin gene (DMD) and from the patient’s brother with a normal dystrophin gene as a healthy control ( Fig. 1A ). Deletion of exon 44 (ΔEx44) disrupts the open reading frame of dystrophin by causing splicing of exon 43 to exon 45 and introducing a premature termination codon ( Fig. 1B ). The reading frame can be restored by using CRISPR-Cas9 gene editing to skip exon 43, which allows splicing between exons 42 and 45, or to skip exon 45, which allows splicing between exons 43 and 46. Alternatively, reframing of exon 43 or 45 can restore the protein reading frame by inserting one nucleotide (+3n+1 insertion) or deleting two nucleotides (+3n−2 deletion).

DISCUSSION

Our results establish a new mouse model of DMD lacking exon 44 of the dystrophin gene, representing one of the most prevalent hotspot regions for dystrophin gene mutations in humans. Correction of exon 44 deletions through exon skipping or reframing of surrounding exons could potentially treat ~12% of patients with DMD. These ΔEx44 DMD mice display the hallmarks of DMD, including myocyte degeneration, regeneration, fibrosis, and fatty infiltration of muscle, as well as loss of contractile function, and will provide a platform for testing and optimizing gene editing strategies and other therapies. The dystrophin exon 44 deletion in these mice and the strategy for restoration of dystrophin expression by skipping exon 45 are analogous to the correction strategy using the oligonucleotide casimersen (SRP-4045), developed by Sarepta, which is designed to restore dystrophin expression in patients with exon 44 deletions by masking the splice acceptor site on exon 45. In a recent clinical trial, eteplirsen, an oligonucleotide that allows exon 51 skipping in patients lacking exon 50, was reported to allow the expression of ~0.5% of the normal level of dystrophin, as measured in biopsy samples from treated patients with DMD after approximately 1 year of continuous treatment (6). By comparison, we observed ~90% restoration of dystrophin protein expression in all muscles and the heart of mice with exon 44 deletion within 4 weeks of a single systemic dose of gene editing components encoded by AAV9. It has been estimated that only 15 to 30% of normal dystrophin levels could provide therapeutic benefits in patients (22, 23).

We show that the ratio of AAVs encoding sgRNA and Cas9 can have a profound effect on the efficiency of gene correction in vivo. Increasing the ratio of AAV-sgRNA to AAV-Cas9 markedly increases gene correction by single-cut CRISPR. There are several potential explanations to account for these observations: (i) AAV-sgRNA may be limiting in vivo, such that more virus enables greater gene editing. Moreover, because association of Cas9 with sgRNA has been reported to induce a conformational change in Cas9 that potentiates gene editing (24), higher levels of sgRNA may ensure higher Cas9 activity in vivo. (ii) sgRNAs are transcribed by RNA polymerase III and are likely to be confined to the nucleus (25, 26). Cas9 protein, derived from translation of Cas9 mRNA in the cytoplasm, can enter nuclei other than those in which the sgRNA was transcribed. Increasing the level of AAV-sgRNA may allow for a higher percentage of nuclei within myofibers to express the sgRNA, thereby enhancing CRISPR-Cas9 genomic editing. (iii) Depletion of sgRNA may occur over time in vivo, and increasing the abundance of sgRNA may ensure continuous editing in myofibers.

When a constant dosage of AAV-Cas9 was administered with higher amounts of AAV-sgRNA, we observed increased Cas9 protein and mRNA expression. Perhaps, the increase in Cas9 expression with sgRNA dosage and the consequent increase in dystrophin restoration lead to a healthier cellular environment for Cas9 expression. The difference in rescue efficiency at different ratios of AAV-sgRNA and AAV-Cas9 potentially correlates with the number of nuclei edited in each cell. Although both cardiac muscle and skeletal muscle are multinucleated, a single cardiomyocyte contains one to four nuclei on average, but one myofiber may contain hundreds of nuclei. Thus, generating dystrophin-expressing myocytes by editing nuclei in one cardiomyocyte is more efficient than in one myofiber. As a result, when supplied with the same amount of sgRNA, cardiac muscle shows better editing efficacy than skeletal muscle. In addition, on the basis of previous reports, it is likely that AAV9 has better tropism for heart tissue than skeletal muscle (27).

Using a single sgRNA against a sequence within exon 45, we observed a high fraction of single nucleotide insertions immediately adjacent to the DNA cut, and these insertions were most commonly an adenosine, corresponding to the next nucleotide adjacent to the site of the initial double-strand DNA break. We made similar observations with single-cut gene editing of exon 51 in mice and dogs lacking exon 50 (11). Because cutting with Cas9 has a propensity for a single nucleotide 5′ overhang four nucleotides 5′ to the cut site, the presence of a thymidine at this position favors the insertion of an adenosine on the complementary strand during DNA repair (28). This single nucleotide insertion has the potential to restore the open reading frame if the exon is out of frame with the preceding exon by a single nucleotide, as in the case of exons 43 and 45. However, this strategy is less likely to restore the open reading frame if two nucleotides are required to reframe the protein due to the low frequency of two-nucleotide insertions after non-homologous end joining (NHEJ). Nevertheless, deletions that remove the splice acceptor or donor sequence of the out-of-frame exon can restore dystrophin in such cases. Notably, we observed a low frequency of integration of AAV ITR sequences at the site of Cas9 cutting in vivo, as observed by previous reports (29, 30).

Our results highlight the effectiveness of single-cut CRISPR gene editing for efficient restoration of dystrophin in vivo. While several studies have also shown that the use of two sgRNAs to mediate Cas9 cutting at distal genomic sites can allow for excision of large intervening genomic regions and restoration of dystrophin expression from mutant alleles (31–33), the efficiency of the double-cut approach is low and is associated with unpredictable genomic rearrangements that we have not observed using only a single sgRNA to direct Cas9 cutting. Thus, we believe that the single-cut CRISPR editing approach represents the most viable clinical approach for correction of dystrophin mutations by gene editing. Of course, it also remains to be determined if the marked effects we have observed here in mice can be scaled up to humans with much larger muscles over a longer time frame.

There are several limitations of our study that should be considered. While we have shown marked restoration of dystrophin protein and muscle structure within 4 weeks of AAV delivery, we do not yet know whether these effects will be sustained or, alternatively, may fade over time. Considering that the majority of cardiomyocytes do not turn over, we expect that the benefits of dystrophin restoration in the heart will be lifelong. However, it remains to be determined if there will be gradual turnover of skeletal muscle following delivery of gene editing components by AAV9. In this regard, Wagers and co-workers have reported that AAV9 infects satellite cells in vivo (34), which could provide a sustained reservoir of cells for long-term maintenance of dystrophin expression. However, we and others have not observed efficient AAV infection of satellite cells in vivo (11, 35). Whether this represents technical differences in delivery approaches remains to be determined.

Possible immunological responses to Cas9 or dystrophin also remain to be investigated over the long term. While we have not observed an immune response to AAV or Cas9, nor to dystrophin, in our previous studies (11), it is conceivable that such responses might be seen over longer time periods. Last, we have tested for possible off-target genomic cutting at sites predicted to have highest homology with the sgRNAs used to correct the ΔEx44 deletion but have not observed any off-target cutting above background. There has been little evidence of off-target effects of CRISPR-Cas9 editing in mice, other than one report that was retracted (36).

In summary, the ΔEx44 DMD mice described here, combined with the optimized ssgRNA and AAV vectors for delivery, should facilitate progress toward long-term correction of dystrophin mutations in mice as a prelude to possible clinical translation.