GENOME EDITING PRIORITY CLAIM

The present application claims benefit of priority to U.S. Provisional Application Serial No. 62/547,590, filed on August 18, 2017, the entire contents of which are hereby incorporated by reference in their entireties. FEDERAL FUNDING SUPPORT CLAUSE

This invention was made with government support under grant no. U54 HD 087351 awarded by National Institutes of Health. The government has certain rights in the invention.

SEQUENCE LISTING

The instant application contains a sequence listing which has been submitted electronically in ASCII format and is hereby incorporated by reference in its entirety. Said ASCII copy, created on August 16, 2018, is named EXON_008_01WO_SeqList_ST25.txt and is 1 MB in size. FIELD OF THE DISCLSOURE

The present disclosure relates to the fields of molecular biology, medicine and genetics. More particularly, the disclosure relates to the use of genome editing to correct mutations in actin binding domain 1 of the DMD gene using an exon-deletion approach. BACKGROUND

Dystrophin is a large intracellular protein that stabilizes muscle membranes against forces associated with contraction and stretch by providing a mechanical link between the intracellular actin cytoskeleton and the transmembrane dystrogly can complex. The DMD gene, one of the largest human genes, encodes dystrophin and is comprised of 79 exons on the Xp21 chromosome. Mutations that disrupt the open reading frame of dystrophin lead to Duchenne muscular dystrophy (DMD), a life-limiting, rapidly progressive form of muscular dystrophy. DMD is a X-linked recessive disorder affecting 1 :5,000 boys. Loss of dystrophin destabilizes muscle membranes, allowing excess calcium into cardiac and skeletal muscle cells, resulting in muscle degeneration and necrosis. Internal deletion mutations in the dystrophin gene {DMD) that preserve the amino- and carboxyl-termini of the protein but eliminate various internal rod domains cause Becker muscular dystrophy (BMD), a relatively mild muscle disease.

There remains a need in the art for compositions and methods for treating DMD.

SUMMARY

Thus, in accordance with the present disclosure, there is provided a composition comprising a sequence encoding a first DMD guide RNA (gRNA) targeting a first genomic target sequence; a sequence encoding a second DMD gRNA targeting a second genomic target sequence; a sequence encoding a first promoter, wherein the first promoter drives expression of the sequence encoding the first DMD gRNA; and a sequence encoding a second promoter, wherein the second promoter drives expression of the sequence encoding the second DMD gRNA; wherein the first genomic target sequence is in any one of introns 2-9 of the dystrophin gene; wherein the second genomic target sequence is in any one of introns 2-9 of the dystrophin gene. In some embodiments, the first genomic target sequence is in intron 2 and the second genomic target sequence is in intron 7 of the dystrophin gene. In some embodiments, the first genomic target sequence is in intron 5 and the second genomic target sequence is in intron 7 of the dystrophin gene. In some embodiments, the first genomic target sequence is in intron 2 and the second genomic target sequence is in intron 9 of the dystrophin gene. In some embodiments, the first genomic target sequence is in intron 7 and the second genomic target sequence is in intron 9 of the dystrophin gene. In some embodiments, the first genomic target sequence is located 5' from a wildtype exon, and the second genomic target sequence is located 3' from the wildtype exon. The wildtype exon may be exon 2, 3, 4, 5, 6, 7, 8 or 9 of dystrophin. In some embodiments, the first genomic target sequence is located 5' from an exon comprising a mutation, and the second genomic target sequence is located 3' from the exon comprising a mutation. The exon comprising a mutation may be exon 2, 3, 4, 5, 6, 7, 8 or 9 of dystrophin. In some embodiments, the sequence encoding the first DMD gRNA and the sequence encoding the second DMD gRNA are identical. In some embodiments, the sequence encoding the first DMD gRNA and the sequence encoding the second DMD gRNA are not identical. In some embodiments, the sequence encoding the first DMD guide RNA is any one of SEQ ID NO: 1 to 5. In some embodiments, the sequence encoding the second DMD guide RNA is any one of SEQ ID NO: 1 to 5. In some embodiments, the sequence of the first DMD guide RNA is any one of SEQ ID NO: 6 to 10. In some embodiments, the sequence of the second DMD guide RNA is any one of SEQ ID NO: 6 to 10. In some embodiments, the sequence encoding the first promoter and the sequence encoding the second promoter are identical. In some embodiments, the sequence encoding the first promoter and the sequence encoding the second promoter are not identical. In some embodiments, at least one of the first promoter and the second promoter is a constitutive promoter. In some embodiments, at least one of the first promoter and the second promoter is an inducible promoter. In some embodiments, at least one of the first promoter and the second promoter is a muscle-specific promoter. In some embodiments, at least one of the first promoter and the second promoter is CK8. In some embodiments, at least one of the first promoter and the second promoter is CK8e. In some embodiments, at least one of the first promoter and the second promoter is selected from the group consisting of the U6 promoter, the HI promoter, and the 7SK promoter. In some embodiments, a first vector comprises the sequence encoding the first DMD gRNA and the first promoter, and a second vector comprises the sequence encoding the second DMD gRNA and the second promoter. In some embodiments, at least one of the first vector and the second vector is a non- viral vector. In some embodiments, the non-viral vector is a plasmid. In some embodiments, a liposome or nanoparticle comprises the non-viral vector. In some embodiments, at least one of the first vector and the second vector is a viral vector. In some embodiments, the viral vector is an adeno-associated viral (AAV) vector. In some embodiments, the AAV vector is replication-defective or conditionally replication defective. In some embodiments, the AAV vector is a recombinant AAV vector. In some embodiments, the AAV vector comprises a sequence isolated or derived from an AAV vector of serotype AAV1, AAV2, AAV 3, AAV4, AAV5, AAV6, AAV7, AAV8, AAV9, AAV 10, AAV11, AAVRh.74 or any combination thereof. In some embodiments, a first vector comprises the sequence encoding the first DMD gRNA, the sequence encoding the second DMD gRNA, the sequence encoding the first promoter, and the sequence encoding the second promoter. In some embodiments, the composition further comprises a sequence encoding a nuclease, such as Cas9. In some embodiments, the sequence encoding the Cas9 is isolated or derived from S. aureus. In some embodiments, the composition comprises a pharmaceutically-acceptable carrier.

Also provided is a cell comprising a composition of the disclosure. The cell may be a mammalian cell, such as a murine cell or a human cell. In some embodiments, the cell is an oocyte (e.g., a non-human oocyte). Also provided are compositions comprising a cell of the disclosure, and a genetically engineered mouse comprising a cell of the disclosure. Also provided is a method of treating a subject in need thereof, the method comprising administering to the subject a therapeutically effective amount of a composition of the disclosure. In some embodiments, the composition is administered systemically. In some embodiments, the composition is administered by an intravenous infusion or injection. In some embodiments, the composition is administered locally. In some embodiments, the composition is administered directly to a muscle tissue, such as a tibialis anterior tissue, a quadriceps tissue, a soleus tissue, a diaphragm tissue, or a heart tissue. In some embodiments, the composition is administered by an intramuscular infusion or injection. In some embodiments, the composition is administered by intra-cardiac injection. In some embodiments, the subj ect is a neonate, an infant, a child, a young adult, or an adult. In some embodiments, the subject has muscular dystrophy. In some embodiments, the subject is a genetic carrier for muscular dystrophy. In some embodiments, the subject is male. In some embodiments, the subject is female. In some embodiments, the subject is less than 10 years old, less than 5 years old, or less than 2 years old.

Also provided herein is the use of the composition of the disclosure in the manufacture of a medicament for the treatment of muscular dystrophy.

Also provided herein is a genetically engineered mouse whose genome comprises a deletion of exon 8 and 9 of the dystrophin gene resulting in an out of frame shift and a premature stop codon in exon 10.

Also provided herein is a genetically engineered mouse produced by a method comprising the steps of: (a) contacting a fertilized oocyte with (i) a Cas9, a first gRNA, and a second gRNA, or (ii) one or more sequences encoding the same, thereby creating a modified oocyte, wherein the first gRNA targets an intron located 5 ' to exon 8 of the dystrophin gene, wherein the second gRNA targets an intron located 3 ' to exon 9 of the dystrophin gene, wherein the contacting causes exons 8 and 9 to be deleted in the modified oocyte, wherein deletion of exons 8 and 9 results in an out of frame shift and a premature stop codon in exon 10; and (b) transferring the modified oocyte into a recipient female. The disclosure also provides a mouse produced by this method.

In some embodiments, the disclosure provides a method of editing an Actin Binding Domain 1 (ABD-1) dystrophin gene defect in a subject comprising contacting a cell with one or more expression constructs expressing Cas9, a first guide RNA and a second guide RNA, wherein the first guide RNA targets a dystrophin intron 5 ' to the gene defect, and the second guide RNA targets a dystrophin intron 3 ' to the gene defect, thereby resulting in an edited dystrophin gene lacking dystrophin exons 3-9. The cell may be a muscle cell, a satellite cell, or an iPSC/iPSC-derived CM. In some embodiments, the Cas9 expression construct is distinct from the expression construct that expresses the first and/or second guide RNAs. In some embodiments, the Cas9 expression construct is the same expression construct as that expressing the first and/or second guide RNAs. In some embodiments, the expression construct(s) is/are a viral vector. In some embodiments, the expression construct(s) is/are a non-viral vector. In some embodiments, the expression construct(s) is/are naked plasmid DNA or chemically- modified mRNA. In some embodiments, (a) the first guide RNA targets dystrophin intron 2 and the second guide RNA targets dystrophin intron 9; (b) the first guide RNA targets dystrophin intron 2, and the second guide RNA targets dystrophin intron 7; (c) the first guide RNA targets dystrophin intron 7, and the second guide RNA targets dystrophin intron 9. In some embodiments, the expression construct(s) is/are provided to the cell in one or more nanoparticles. In some embodiments, the viral vector is an AAV vector, such as AAV-9. In some embodiments, the contacting comprises administration of AAV vector to the subject, such as by intra-muscular, intra-peritoneal (IP), retro-orbital (RO), or intra-cardiac injection. In some embodiments, the expression construct(s) is/are delivered to and iPSC/iPSC-derived CM or directly to a muscle tissue, such as tibialis anterior, quadriceps, soleus, diaphragm or heart. In some embodiments, the expression construct(s) is/are delivered systemically. In some embodiments, the subject exhibits normal dystrophin-positive myofibers and/or mosaic dystrophin-positive myofibers containing centralized nuclei. In some embodiments, the subject exhibits a decreased serum CK level as compared to a serum CK level prior to contacting. In some embodiments, the subject exhibits improved grip strength as compared to a serum CK level prior to contacting. In some embodiments, the first guide RNA is encoded by the DNA sequence 5'-AATTAATCTGCCGAAGATGA-3' (SEQ ID NO: 1). In some embodiments, the second guide RNA is encoded by the DNA sequence 5'- AAACAAACCAGCTCTTCACG-3' (SEQ ID NO: 5). In some embodiments, the expression construct(s) is/are delivered to a human iPS cell by nucleofection. In some embodiments, the method comprises further comprising identifying an ABD-1 target based on reference to a Duchenne mutation database. In some embodiments, the first and second guide RNAs are encoded by and expressed from the same expression construct, or from distinct expression constructs. In some embodiments, one or more promoters in the expression construct(s) is/are RNA polymerase III promoters. In some embodiments, the mutant dystrophin exon is exon 3, 4, 5, 6, 7, 8 or 9. Other objects, features and advantages of the present disclosure will become apparent from the following detailed description. It should be understood, however, that the detailed description and the specific examples, while indicating preferred embodiments, are given by way of illustration only, since various changes and modifications within the spirit and scope of the disclosure will become apparent to those skilled in the art from this detailed description.

BRIEF DESCRIPTION OF THE DRAWINGS

The following drawings form part of the present specification and are included to further demonstrate certain aspects of the present disclosure. The disclosure may be better understood by reference to one or more of these drawings in combination with the detailed description of specific embodiments presented herein.

FIG. 1A-F. Generating an iDMD model by deleting DMD exons 8-9 using CRISPR/Cas9-mediated genome editing (FIG 1A) Strategy showing CRISPR/Cas9- mediated genomic editing of wild type (WT) DMD to generate ΔΕχ8-9 iDMD. Shape and color of boxes denoting DMD exons indicate reading frame and protein coding domains. Yellow designates actin binding domain-1 (ABD-1). Blue marks part of the central rod domain. Red lines indicate actin binding sites (ABSl, ABS2 and ABS3). Arrowheads mark targeting site of guide RNAs (gRNAs). Stop sign marks exon with stop codon. (FIG. IB) Sequences of gRNAs (SEQ ID NO: 93, SEQ ID NO: 5) and their targeting sites within intron 7 (top, SEQ ID NO: 94-45) and intron 9 (bottom, SEQ ID NO: 96-97). gRNAs were designed to target 3' region of intron 7 (gRNA-7) and 5' region of intron 9 (gRNA-9). PAM sites are highlighted in red. (FIG. 1C) PCR genotyping of control and ΔΕχ8-9 iDMD induced pluripotent stem cell (iPSC) lines using primers upstream and downstream of the gRNA targeting sites (top) and within intron 7 flanking the gRNA-7 targeting site (bottom). Sequencing of PCR product of ΔΕχ8-9 iDMD validates splicing of intron 7 to intron 9 (SEQ ID NO: 98). PCR primers are indicated by arrows. Arrowhead indicates gRNA targeting site. M denotes marker lane. (FIG. ID) RT-PCR analysis of dystrophin mRNA expression in control and ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes. Forward primer targeting exon 1 and reverse primer targeting exon 10 were used. Sequencing confirmed splicing of exon 7 to exon 10, introducing a stop-codon (SEQ ID NO: 99-100). (FIG. IE) Western blot analysis showing dystrophin protein expression in iPSC- derived cardiomyocytes using anti-dystrophin antibody. Vinculin was used as loading control. n=7 for each group. (FIG. IF) Immunocytochemistry representations of iPSC-derived cardiomyocytes with anti-dystrophin (red) and anti-troponin I (green). Nuclei are stained with Hoechst 33342 (blue). Scale bar = 50 microns. n=4 for each group. FIG. 2A-F. Correcting AEx8-9 iDMD by exon deletion of exons 3-9 to restore dystrophin expression. (FIG. 2A) Three strategies were used to correct ΔΕχ8-9 iDMD by CRISPR/Cas9-mediated genomic editing: (1) deleting exons 3-7 to generate ΔΕχ3-9, (2) deleting exons 6-7 to generate ΔΕχ6-9, and (3) deleting exons 7-11 to generate ΔΕχ7-11. Shape and color of boxes denoting DMD exons indicate reading frame and protein coding domains. Yellow designates actin binding domain-1 (ABD-1). Blue marks part of the central rod domain. Red lines indicate actin binding sites (ABS 1, ABS2 and ABS3). Red exon indicates exon with stop codon. (FIG. 2B) Illustration showing deletion of exons 3-7 to generate ΔΕχ3-9. Sequences of gRNAs (SEQ ID NO: 1, SEQ ID NO: 4) and their targeting sites within intron 2 (top, SEQ ID NO: 101-102)) and intron 7 (bottom, SEQ ID NO: 104-105). gRNAs were designed to target 3' region of intron 2 (gRNA-2) and 5' region of intron 7 (gRNA-7). Arrowheads mark targeting site of gRNAs. PAM sites are highlighted in red. (FIG. 2C) PCR genotyping of control, ΔΕχ8-9 iDMD and two clones of ΔΕχ3-9 induced pluripotent cell (iPSC) lines using primers upstream and downstream of the gRNA targeting sites (top) and within intron 2 flanking the gRNA-2 targeting site (bottom). Sequencing of PCR product of ΔΕχ3-9 validates splicing of intron 2 to intron 7 (SEQ ID NO: 103). PCR primers are indicated by arrows. Arrowhead indicates gRNA targeting site. M denotes marker lane. (FIG. 2D) RT- PCR analysis of dystrophin mRNA expression in control, ΔΕχ8-9 iDMD, and two clones of ΔΕχ3-9 iPSC-derived cardiomyocytes. Forward primer targeting exon 1 and reverse primer targeting exon 10 were used. Sequencing confirmed splicing of exon 2 to exon 10 restoring the open reading frame (SEQ ID NO: 106-107). a-actinin was used as loading control. (FIG. 2E) Western blot analysis showing dystrophin protein expression in iPSC-derived cardiomyocytes using anti-dystrophin antibody. Vinculin was used as loading control. n=7 for control and ΔΕχ8-9 iDMD, n=3 for ΔΕχ3-9 clone 1 and ΔΕχ3-9 clone 2 n=7 for control and ΔΕχ8-9 iDMD, n=3 for ΔΕχ3-9 clone 1 and ΔΕχ3-9 clone 2. (FIG. 2F) Immunocytochemistry representations of iPSC-derived cardiomyocytes with anti-dystrophin (red) and anti-troponin I (green). Nuclei are stained with Hoechst 33342 (blue). Scale bar = 50 microns. n=4 for control and ΔΕχ8-9 iDMD, n=2 for ΔΕχ3-9 clone 1, and n=l for ΔΕχ3-9 clone 2.

FIG. 3A-E. Correcting AEx8-9 iDMD by deleting exons 6 and 7 to restore dystrophin protein expression. (FIG. 3A) Illustration showing deletion of exons 6-7 to generate ΔΕχ6-9. Sequences of guide RNAs (gRNAs) (SEQ ID NO: 2-3) and their targeting sites within intron 5 (top, SEQ ID NO: 108-109) and intron 7 (bottom, SEQ ID NO: 110-111). gRNAs were designed to target 3' region of intron 5 (gRNA-5) and 5' region of intron 7 (gRNA-7). Arrowheads mark targeting site of gRNAs. PAM sites are highlighted in red. (FIG. 3B) PCR genotyping of control, ΔΕχ8-9 iDMD and two clones of ΔΕχ6-9 induced pluripotent stem cell (iPSC) lines using primers upstream and downstream of the gRNA targeting sites (top) and within intron 5 flanking the gRNA-5 targeting site (bottom). Sequencing of PCR product of ΔΕχ6-9 validates splicing of intron 5 to intron 7 (SEQ ID NO: 112). PCR primers are indicated by arrows. Arrowhead indicates gRNA targeting site. M denotes marker lane. (FIG. 3C) RT-PCR analysis of dystrophin mRNA expression in control, ΔΕχ8-9 iDMD, and two clones of ΔΕχ6-9 iPSC-derived cardiomyocytes. Forward primer targeting exon 5 and reverse primer targeting exon 10 were used. Sequencing confirmed splicing of exon 5 to exon 10 restoring the open reading frame (SEQ ID NO: 113-114). a-actinin was used as loading control. (FIG. 3D) Western blot analysis showing dystrophin protein expression in iPSC- derived cardiomyocytes using anti-dystrophin antibody. Vinculin was used as loading control. n=7 for control and ΔΕχ8-9 iDMD, n=3 for ΔΕχ6-9 clone 1 and ΔΕχ6-9 clone 2. (FIG. 3E) Immunocytochemistry representations of iPSC-derived cardiomyocytes with anti-dystrophin (red) and anti-troponin I (green). Nuclei are stained with Hoechst 33342 (blue). Scale bar = 50 microns. n=4 for control and ΔΕχ8-9 iDMD, n=3 for ΔΕχ6-9 clone 1 and ΔΕχ6-9 clone 2.

FIG. 4A-F. Correcting AEx8-9 iDMD by deleting exons 7-11 partially restores dystrophin protein expression. (FIG. 4A) Illustration showing deletion of exons 7-11 to generate ΔΕχ7-11. Sequences of guide RNAs (gRNAs) (SEQ ID NO: 115, SEQ ID NO: 118) and their targeting sites within intron 6 (top, SEQ ID NO: 116-117) and intron 11 (bottom, SEQ ID NO: 118-119). gRNAs were designed to target 3' region of intron 6 (gRNA-6) and 5' region of intron 11 (gRNA-11). Arrowheads mark gRNA targeting site. PAM sites are highlighted in red. (FIG. 4B) PCR genotyping of control, ΔΕχ8-9 iDMD and two clones of ΔΕχ7-11 induced pluripotent stem cell (iPSC) lines using primers upstream and downstream of the gRNA targeting sites (top) and within intron 6 flanking the gRNA-6 targeting site (bottom). Sequencing of PCR product of ΔΕχ7-11 validates splicing of intron 6 to intron 11 (SEQ ID NO: 121). PCR primers are indicated by arrows. Arrowhead indicates gRNA targeting site. M denotes marker lane. (FIG. 4C) RT-PCR analysis of dystrophin mRNA expression in control, ΔΕχ8-9 iDMD, and two clones of ΔΕχ7-11 iPSC-derived cardiomyocytes. Forward primer targeting exon 5 and reverse primer targeting exon 12 were used. Sequencing confirmed splicing of exon 6 to exon 12 restoring the open reading frame (SEQ ID NO: 122-123). a- actinin was used as loading control. (FIG. 4D) Western blot analysis showing dystrophin protein expression in iPSC-derived cardiomyocytes using anti-dystrophin antibody. Vinculin was used as loading control. n=7 for control and ΔΕχ8-9 iDMD, n=3 for ΔΕχ7-11 clone 1 and ΔΕχ7-11 clone 2. (FIG. 4E) Immunocytochemistry representations of iPSC-derived cardiomyocytes with anti-dystrophin (red) and anti-troponin I (green). Nuclei are stained with Hoechst 33342 (blue). Scale bar = 50 microns. n=4 for control and ΔΕχ8-9 iDMD, n=2 for ΔΕχ7-11 clone 1 and n=l for ΔΕχ7-11 clone 2. (FIG. 4F) Western blot analysis of ΔΕχ7-11 iPSC-derived cardiomyocytes treated with proteasome inhibitor MG132 for 60 hours using anti-dystrophin antibody. GAPDH was used as loading control. n=2.

FIG 5A-G. Functional analysis of iPSC-derived cardiomyocytes. (FIG. 5A) Representative recordings of spontaneous Ca ²⁺ activity of induced pluripotent cell (iPSC)- derived cardiomyocytes loaded with Ca ²⁺ indicator Fluo-4AM. Traces show change in fluorescence intensity (F) in relationship to resting fluorescence intensity (F ₀ ). (FIG. 5B) Relative time to peak (TTP), (FIG. 5C) decay (tau) and (FIG. 5D) transient duration (TD), as measured by calcium imaging. (FIG. 5E) Arrhythmic iPSC-derived cardiomyocytes were identified based on calcium activity. For all corrected iPSC-derived cardiomyocyte lines (panels B-E), data were obtained from two independent clones. n=113 for control; n=105 for ΔΕχ8-9 iDMD; n=129 for ΔΕχ3-9; n=121 for ΔΕχ6-9; and n=122 for ΔΕχ7-11 iPSC-derived cardiomyocyte lines from 5-11 independent experiments. Data are represented as mean ± s.e.m. *P < 0.05 by one-way ANOVA. (FIG. 5F) Force of EHM contraction in the presence of increasing Ca ²⁺ concentration. EHM generated from two independent control lines (n=8), ΔΕχ8-9 iDMD (n=6), ΔΕχ3-9 (n=14), ΔΕχ6-9 (n=3), and ΔΕχ7-11 (n=3). (FIG. 5G) Representative recordings of EHM contractions from the indicated groups (same EHM as in FIG. 5F). Data are represented as mean ± s.e.m. *P < 0.05 by two-way ANOVA and Tukey's post-hoc test.

FIG. 6A-I Correction of DMD patient-derived iPSCs by deleting exons 8 and 9. (FIG. 6A) Strategy showing CRISPR/Cas9-mediated genomic editing of DMD (Duchene muscular dystrophy) patient (ρΔΕχ3-7) to generate corrected ρΔΕχ3-9 induced pluripotent cell (iPSC) line. Shape and color of boxes denoting DMD exons indicate reading frame and protein coding domains. Yellow designates ABD-1. Blue marks part of central rod domain. Red lines indicate ABS1. Arrowheads mark targeting site of guide RNAs (gRNAs). Red box marks exon with stop codon. Sequences of gRNAs (SEQ ID NO: 124, SEQ ID NO: 5) and their targeting sites within intron 7 (top, SEQ ID NO: 125-126) and intron 9 (bottom, SEQ ID NO: 128-129). gRNAs were designed to target 3' region of intron 7 (gRNA-7) and 5' region of intron 9 (gRNA-9). PAM sites are highlighted in red. (FIG. 6B) PCR genotyping of pAEx3-7and ρΔΕχ3-9 iPSC lines using primers upstream and downstream of the gRNA targeting sites. Sequencing of PCR product of ρΔΕχ3-9 validates splicing of intron 7 to intron 9 (SEQ ID NO: 127). PCR primers are indicated by arrows. Arrowheads mark targeting site of gRNAs. M denotes marker lane. (FIG. 6C) RT-PCR analysis of dystrophin mRNA expression in control, ρΔΕχ3-7, ρΔΕχ3-9 iPSC-derived cardiomyocytes. Forward primer targeting 5'UTR and reverse primer targeting exon 10 were used. Sequencing of ρΔΕχ3-7 confirmed splicing of exon 2 to exon 8 (SEQ ID NO: 130-131), introducing a stop-codon. Sequencing ρΔΕχ3-9 confirmed splicing of exon 2 to exon 10 (SEQ ID NO: 132-133) restoring the open reading frame, a-actinin was used as loading control. (FIG. 6D) Western blot analysis showing dystrophin protein expression in iPSC-derived cardiomyocytes using anti-dy strophin antibody. Vinculin was used as loading control. n=7 for control, n=3 for ρΔΕχ3-7 and n=3 for ρΔΕχ3-9. (FIG. 6E) Immunocytochemistry representations of iPSC-derived cardiomyocytes with anti- dystrophin (red) and anti-troponin I (green). Nuclei are stained with Hoechst 33342 (blue). Scale bar = 50 microns. n=4 for control, n=2 for ρΔΕχ3-7 and n=2 for ρΔΕχ3-9. (FIG. 6F) Relative time to peak (TTP), (FIG. 6G) decay (tau), (FIG. 6H) transient duration (TD) and (FIG. 61) percent of arrhythmic cells were measured based on calcium activity of control (n=45), DMD patient ρΔΕχ3-7 (n=40) and corrected ρΔΕχ3-9 (n=65) iPSC-derived cardiomyocytes from 3-5 independent experiments. Data are represented as mean ± s.e.m. *P < 0.05 by one-way ANOVA (FIGS. 6F-H).

FIG. 7A-C. Dystrophin protein expression level. Related to FIGS 3A-E and FIGS

4A-F. (FIG. 7A) Dystrophin expression levels in ΔΕχ8-9 iDMD (n=8), ΔΕχ3-9 (n=4), ΔΕχ6- 9 (n=6), and ΔΕχ7-11 (n=5) iPSC-derived cardiomyocytes compared to control cell line, indicated by dashed line. Data are represented as mean ± s.e.m. *P < 0.05. (FIG. 7B) Western blot analysis of dystrophin (top), Myosin heavy chain, Myh, (middle) and GAPDH (bottom) expression in control, ΔΕχ8-9 iDMD, ΔΕχ3-9, ΔΕχ6-9, and ΔΕχ7-11 iPSC-derived cardiomyocytes. Arrowhead indicates residual dystrophin protein after stripping the blot. Arrow indicates myosin heavy chain expression. (FIG. 7C) Western blot analysis of ΔΕχ7-11 clone 2 iPSC-derived cardiomyocytes treated with proteasome inhibitor MG132 for 60 hours using anti-dystrophin antibody. GAPDH and Vinculin were used as loading control.

FIG. 8A-C. Generation and functional analysis of engineered heart muscle

(EHM). Related to FIGS. 5A-G. (FIG. 8A) Schematic diagram of EHM generation. (FIG. 8B) Percentage of a-actinin-positive (ACTN2+) cardiomyocytes before EHM generation. n=7 total EHM analyzed. (FIG. 8C) Percentage of EHM arrhythmic contractions. FIG. 9A-E. Generation of DMD mouse model by deletion of Dmd exons 8 and 9.

(FIG. 9A) Outline of the CRISPR/Cas9 strategy used to generate the DMD mouse model, ΔΕχ8-9 by excising exons 8 and 9. (FIG. 9B) Hematoxylin and eosin (H&E) immunostaining of quadriceps, diaphragm and heart in wild type (WT) and ΔΕχ8-9 DMD mice. (FIG. 9C) Dystrophin immunostaining of quadriceps, diaphragm and heart of wild type (WT) and ΔΕχ8- 9 DMD mice. Dystrophin immunostains in red. Nucleus are marked by DAPI stain in blue. (FIG. 9D) Grip strength measured in WT and ΔΕχ8-9 DMD mice. (FIG. 9E) Serum creatine kinase (CK), a marker of muscle dystrophy that reflects muscle damage and membrane leakage was measured in WT and ΔΕχ8-9 DMD mice.

SEQUENCE TABLES

TABLE 1. Human DMD guide RNA for actin binding domain region

TABLE 2. Mouse DMD guide RNA for actin binding domain region

DETAILED DESCRIPTION

DMD is associated with greater than 4,000 mutations in the DMD gene, which are comprised primarily of exon deletions, as well as exon duplications and small mutations that include point mutations. DMD mutations cluster into two hot spot regions of the gene. One hot spot is located within the 5' region of the gene, encompassing exons 2-20. These mutations account for -15% of all exon deletions and -50% of all exon duplications within the DMD gene. Deletion of exons 3-7 are the most frequent. The second DMD hot spot is located in the distal region of the DMD gene, between exons 45-55, accounting for -70% of all exon deletions and -15% of all exon duplications. Because a majority of patient mutations carry deletions in these hotspots, a therapeutic approach for skipping certain exon applies to large group of patients. The rationale of the exon skipping approach is based on the genetic difference between DMD and Becker muscular dystrophy (BMD) patients. In DMD patients, the reading frame of dystrophin mRNA is disrupted resulting in prematurely truncated, non-functional dystrophin proteins. BMD patients have mutations in the DMD gene that maintain the reading frame allowing the production of internally deleted, but partially functional dystrophins leading to much milder disease symptoms compared to DMD patients.

Dystrophin protein consists of 3,685 amino acids and can be separated into four domains: 1) the actin binding domain (ABD-1), which consists of amino acids 14-240 and connects the filamentous elements of the cytoskeleton to the cell membrane; 2) the central rod domain, composed of 24 spectrin-like repeats and the second actin-binding domain (ABD-2); 3) the cysteine-rich domain; and 4) the carboxyl-terminal domain. Together, these four domains provide the function of dystrophin as a structural link between the cytoskeleton and extracellular matrix to maintain muscle integrity. The dystrophin protein can tolerate internal deletions that maintain a subset of the rod domains with intact amino- and carboxyl-termini regions, resulting in mild loss of muscle function, as seen in patients with BMD. There have been numerous strategies to convert DMD to BMD by skipping or deleting out-of-frame exons and restoring the expression of truncated forms of dystrophin protein. Similarly, shortened forms of dystrophin, referred to as mini- or micro-dystrophins, are being developed for gene therapy. It should be noted that the precise consequences of in-frame deletions on the stability and function of dystrophin are not predictable a priori, as some in-frame deletions cause severe disease while others have only mild effects. Thus, it is important to analyze the dystrophin protein products generated from in-frame deletions before reaching conclusions regarding their potential therapeutic effects.

Previously, the inventors and others used CRISPR/Cas9-mediated genome editing to permanently correct dystrophin mutations in mouse models of DMD and patient-derived muscle cells. These efforts focused mainly on correcting mutations in the spectrin-like repeat region to restore dystrophin function by generating truncated dystrophin protein, similar to the forms associated with BMD. The ABD-1 of dystrophin contains three actin-binding sites (ABS1-3) that associate with F-actin and are essential for the stabilization of muscle membranes by dystrophin. Little emphasis has been put on editing the ABD-1 region of the DMD gene, although -7% of DMD patients have mutations in the ABD-1 domain. In fact, exons 2-7, which encode part of the ABD-1, are the most frequently mutated portion of the 5'- proximal hot spot. In-frame deletions and missense mutations of the 5' region of the DMD gene that affect the ABD-1 structure have been associated with a decrease in dystrophin protein stability, reduced actin binding affinity, and protein mis-folding and degradation, suggesting that restoring the open reading frame of ABD-1 mutations by genome editing strategies may not be sufficient to correct DMD. On the other hand, medical case studies have reported patients with deletions in exons 3-9 in the DMD gene that exhibit no apparent phenotype, suggesting that precise deletion of exons 3-9 may be an effective approach to correct mutations in the ABD-1 region. Accordingly, uncertainty remains regarding whether gene editing in this region is or is not corrective.

As described in further detail below, the instant inventors demonstrate that gene editing in the actin-binding domain region of the dystrophin gene is a viable method for treating DMD.

Human induced pluripotent stem cells (iPSCs) offer a unique opportunity for studying human diseases in a dish. Their unlimited proliferation, ability to undergo clonal selection, and differentiation into different cell types provide a reliable platform for testing gene editing strategies to either create or correct human mutations. Differentiation of DMD-derived iPSCs to cardiomyocytes prior to and following genome editing allows the analysis of DMD phenotypes by assessing dystrophin expression and cardiomyocyte function, such as muscle contractility and Ca ²⁺ handling. Assembly of iPSC-derived cardiomyocytes as engineered heart muscle (EHM) allows for direct and controlled measurements of heart muscle force of contraction.

Here the inventors introduced an exon 8-9 deletion (ΔΕχ8-9) in the DMD gene of healthy donor-derived iPSCs to generate a DMD iPSC line. This ΔΕχ8-9 DMD iPSC line allowed correction of the DMD mutation by various gene editing strategies and then assess truncated dystrophin functionality in comparison to isogenic control cells. They corrected the exon 8-9 deletion mutation using three different exon-deletion strategies to restore the open reading frames: 1) deleting exons 3-7 to generate ΔΕχ3-9, which excises the ABS-2 and ABS- 3 regions; 2) deleting exons 6-7 to generate ΔΕχ6-9, which excises ABS-3; and 3) deleting exons 7-11 to generate ΔΕχ7-11, which leaves all three ABS regions intact. The inventors show that deletion DMD exons 3-9 generates a truncated dystrophin but maintains the structure of dystrophin such that it restores cardiomyocyte functionality. These findings provide a promising strategy for correction of DMD mutations within the proximal hot spot of the DMD gene by genomic editing, allowing restoration of dystrophin expression and function. These and other aspects of the disclosure are reproduced below.

To further assess the efficiency and optimize CRISPR/Cas9-mediated exon skipping in vivo, a mimic of the human ABD-1 region mutation was generated in a mouse model by deleting the exons 8 and 9 using CRISPR/Cas9 system directed by two single guide RNAs (sgRNAs). The ΔΕχ8-9 DMD mouse model exhibits dystrophic myofibers, increased serum creatine kinase level, and reduced muscle function, thus providing a new representative model of DMD. These and other aspects of the disclosure are reproduced below.

As used herein the specification, "a" or "an" may mean one or more. As used herein in the claim(s), when used in conjunction with the word "comprising", the words "a" or "an" may mean one or more than one.

The use of the term "or" in the claims is used to mean "and/or" unless explicitly indicated to refer to alternatives only or the alternatives are mutually exclusive, although the disclosure supports a definition that refers to only alternatives and "and/or." As used herein "another" may mean at least a second or more. Throughout this application, the term "about" is used to indicate that a value includes the inherent variation of error for the device, for the method being employed to determine the value, or that exists among the study subjects. Such an inherent variation may be a variation of ±10% of the stated value.

I. Duchenne Muscular Dystrophy

A. Background

Duchenne muscular dystrophy (DMD) is a recessive X-linked form of muscular dystrophy, affecting around 1 in 5000 boys, which results in muscle degeneration and premature death. The disorder is caused by a mutation in the gene dystrophin (see GenBank Accession No. NC_000023.11), located on the human X chromosome, which codes for the protein dystrophin (GenBank Accession No. AAA53189; SEQ ID NO. 15), the sequence of which is reproduced below:

1 MLWWEEVEDC YEREDVQKKT FTKWVNAQFS KFGKQHIENL FSDLQDGRRL LDLLEGLTGQ

61 KLPKEKGSTR VHALNNVNKA LRVLQNNNVD LVNIGSTDIV DGNHKLTLGL IWNIILHWQV

121 KNVMKNIMAG LQQTNSEKIL LSWVRQSTRN YPQVNVINFT TSWSDGLALN ALIHSHRPDL

181 FDWNSWCQQ SATQRLEHAF NIARYQLGIE KLLDPEDVDT TYPDKKSILM YITSLFQVLP

241 QQVSIEAIQE VEMLPRPPKV TKEEHFQLHH QMHYSQQITV SLAQGYERTS SPKPRFKSYA

301 YTQAAYVTTS DPTRSPFPSQ HLEAPEDKSF GSSLMESEV LDRYQTALEE VLSWLLSAED

361 TLQAQGEISN DVEWKDQFH THEGYMMDLT AHQGRVGNIL QLGSKLIGTG KLSEDEETEV

421 QEQMNLLNSR WECLRVASME KQSNLHRVLM DLQNQKLKEL NDWLTKTEER TRKMEEEPLG

481 PDLEDLKRQV QQHKVLQEDL EQEQVRVNSL THMWWDES SGDHATAALE EQLKVLGDRW

541 ANICRWTEDR WVLLQDILLK WQRLTEEQCL FSAWLSEKED AVNKIHTTGF KDQNEMLSSL

601 QKLAVLKADL EKKKQSMGKL YSLKQDLLST LKNKSVTQKT EAWLDNFARC WDNLVQKLEK

661 STAQISQAVT TTQPSLTQTT VMETVTTVTT REQILVKHAQ EELPPPPPQK KRQITVDSEI

721 RKRLDVDITE LHSWITRSEA VLQSPEFAI F RKEGNFSDLK EKV AIEREK AEKFRKLQDA

781 SRSAQALVEQ MV EGV ADS IKQASEQLNS RWIEFCQLLS ERLNWLEYQN NIIAFYNQLQ

841 QLEQMTTTAE NWLKIQPTTP SEPTAIKSQL KICKDEV RL SGLQPQIERL KIQSIALKEK

901 GQGPMFLDAD FVAFTNHFKQ VFSDVQAREK ELQTIFDTLP PMRYQETMSA IRTWVQQSET

961 KLSIPQLSVT DYEIMEQRLG ELQALQSSLQ EQQSGLYYLS TTVKEMSKKA PSEISRKYQS

1021 EFEEIEGRWK KLSSQLVEHC QKLEEQMNKL RKIQNHIQTL KKWMAEVDVF LKEEWPALGD

1081 SEILKKQLKQ CRLLVSDIQT IQPSLNSV E GGQKIKNEAE PEFASRLETE LKELNTQWDH

1141 MCQQVYARKE ALKGGLEKTV SLQKDLSEMH EWMTQAEEEY LERDFEYKTP DELQKAVEEM

1201 KRAKEEAQQK EAKVKLLTES VNSVIAQAPP VAQEALKKEL ETLTTNYQWL CTRLNGKCKT

1261 LEEVWACWHE LLSYLEKANK WLNEVEFKLK TTENIPGGAE EI SEVLDSLE NLMRHSEDNP

1321 NQIRILAQTL TDGGVMDELI NEELETFNSR WRELHEEAVR RQKLLEQSIQ SAQETEKSLH

1381 LIQESLTFID KQLAAYIADK VDAAQMPQEA QKIQSDLTSH EI SLEEMKKH NQGKEAAQRV

1441 LSQIDVAQKK LQDVSMKFRL FQKPANFELR LQESKMILDE VKMHLPALET KSVEQEWQS 1501 QLNHCVNLYK SLSEVKSEVE MVIKTGRQIV QKKQTENPKE LDERVTALKL HYNELGAKVT

1561 ERKQQLEKCL KLSRKMRKEM NVLTEWLAAT DMELTKRSAV EGMPSNLDSE VAWGKATQKE

1621 IEKQKVHLKS ITEVGEALKT VLGKKETLVE DKLSLLNSNW IAVTSPAEEW LNLLLEYQKH

1681 METFDQNVDH ITKWIIQADT LLDESEKKKP QQKEDVLKRL KAELNDIRPK VDSTRDQAAN

1741 LMANRGDHCR KLVEPQI SEL NHRFAAI SHR IKTGKASI PL KELEQFNSDI QKLLEPLEAE

1801 IQQGVNLKEE DFNKDMNEDN EGTVKELLQR GDNLQQRITD ERKREEIKIK QQLLQTKHNA

1861 LKDLRSQRRK KALEI SHQWY QYKRQADDLL KCLDDIEKKL ASLPEPRDER KIKEIDRELQ

1921 KKKEELNAVR RQAEGLSEDG AAMAVEPTQI QLSKRWREIE SKFAQFRRLN FAQIHTVREE

1981 TMMVMTEDMP LEISYVPSTY LTEITHVSQA LLEVEQLLNA PDLCAKDFED LFKQEESLKN

2041 IKDSLQQSSG RIDIIHSKKT AALQSATPVE RVKLQEALSQ LDFQWEKVNK MYKDRQGRFD

2101 RSVEKWRRFH YDIKIFNQWL TEAEQFLRKT QI PENWEHAK YKWYLKELQD GIGQRQTWR

2161 TLNATGEEII QQSSKTDASI LQEKLGSLNL RWQEVCKQLS DRKKRLEEQK NILSEFQRDL

2221 NEFVLWLEEA DNIASIPLEP GKEQQLKEKL EQVKLLVEEL PLRQGILKQL NETGGPVLVS

2281 APISPEEQDK LENKLKQTNL QWIKVSRALP EKQGEIEAQI KDLGQLEKKL EDLEEQLNHL

2341 LLWLSPIRNQ LEIYNQPNQE GPFDVQETEI AVQAKQPDVE EILSKGQHLY KEKPATQPVK

2401 RKLEDLSSEW KAVNRLLQEL PAKQPDLAPG LTTIGASPTQ TVTLVTQPW TKETAISKLE

2461 MPSSLMLEVP ALADFNRAWT ELTDWLSLLD QVIKSQRVMV GDLEDINEMI IKQKATMQDL

2521 EQRRPQLEEL ITAAQNLKNK TSNQEARTI I TDRIERIQNQ WDEVQEHLQN RRQQLNEMLK

2581 DSTQWLEAKE EAEQVLGQAR AKLESWKEGP YTVDAIQKKI TETKQLAKDL RQWQTNVDVA

2641 NDLALKLLRD YSADDTRKVH MITENINASW RSIHKRVSER EAALEETHRL LQQFPLDLEK

2701 FLAWLTEAET TANVLQDATR KERLLEDSKG VKELMKQWQD LQGEIEAHTD VYHNLDENSQ

2761 KILRSLEGSD DAVLLQRRLD NMNFKWSELR KKSLNIRSHL EASSDQWKRL HLSLQELLVW

2821 LQLKDDELSR QAPIGGDFPA VQKQNDVHRA FKRELKTKEP VIMSTLETVR I FLTEQPLEG

2881 LEKLYQEPRE LPPEERAQNV TRLLRKQAEE VNTEWEKLNL HSADWQRKID ETLERLQELQ

2941 EATDELDLKL RQAEVIKGSW QPVGDLLIDS LQDHLEKVKA LRGEIAPLKE NVSHVNDLAR

3001 QLTTLGIQLS PYNLSTLEDL NTRWKLLQVA VEDRVRQLHE AHRDFGPASQ HFLSTSVQGP

3061 WERAISPNKV PYYINHETQT TCWDHPKMTE LYQSLADLNN VRFSAYRTAM KLRRLQKALC

3121 LDLLSLSAAC DALDQHNLKQ NDQPMDILQI INCLTTIYDR LEQEHNNLVN VPLCVDMCLN

3181 WLLNVYDTGR TGRIRVLSFK TGIISLCKAH LEDKYRYLFK QVASSTGFCD QRRLGLLLHD

3241 SIQIPRQLGE VASFGGSNIE PSVRSCFQFA NNKPEIEAAL FLDWMRLEPQ SMVWLPVLHR

3301 VAAAETAKHQ AKCNICKECP I IGFRYRSLK HFNYDICQSC FFSGRVAKGH KMHYPMVEYC

3361 TPTTSGEDVR DFAKVLKNKF RTKRYFAKHP RMGYLPVQTV LEGDNMETPV TLINFWPVDS

3421 APASSPQLSH DDTHSRIEHY ASRLAEMENS NGSYLNDSIS PNESIDDEHL LIQHYCQSLN

3481 QDSPLSQPRS PAQILISLES EERGELERIL ADLEEENRNL QAEYDRLKQQ HEHKGLSPLP

3541 SPPEMMPTSP QSPRDAELIA EAKLLRQHKG RLEARMQILE DHNKQLESQL HRLRQLLEQP

3601 QAEAKVNGTT VSSPSTSLQR SDSSQPMLLR WGSQTSDSM GEEDLLSPPQ DTSTGLEEVM

3661 EQLNNSFPSS RGRNTPGKPM REDTM In humans, dystrophin mRNA contains 79 exons. Dystrophin mRNA is known to be alternatively spliced, resulting in various isoforms. Exemplary dystrophin isoforms are listed in Table 3.

Table 3: Dystrophin isoforms

Sequence Nucleic Acid Nucleic Protein Accession Protein Description

Name Accession No. Acid No. SEQ ID

SEQ ID NO:

NO:

Dystrophin NM 004009.3 20 NP 004000.1 21 Transcript Variant:

Dp427pl transcript Dp427pl isoform initiates from a unique promoter/exon 1 located in what corresponds to the first intron of transcript Dp427m. The transcript adds the common exon 2 of Dp427m and has a similar length (14 kb). The Dp427pl isoform replaces the

ML WWEEVED C Y - start of Dp427m with a unique N-terminal MSEVSSD aa sequence.

Dystrophin NM 004011.3 22 NP 004002.2 23 Transcript Variant:

Dp260-1 transcript Dp260-1 uses isoform exons 30-79, and originates from a promoter/exon 1 sequence located in intron 29 of the dystrophin gene. As a result, Dp260-1 contains a 95 bp exon 1 encoding a unique N- terminal 16 aa MTEIILLIFFPAYFLN- sequence that replaces amino acids 1-1357 of the full-length dystrophin product (Dp427m isoform).

Dystrophin NM 004012.3 24 NP 004003.1 25 Transcript Variant:

Dp260-2 transcript Dp260-2 uses isoform exons 30-79, starting from a promoter/exon 1 sequence located in intron 29 of the dystrophin gene that is alternatively spliced and lacks N-terminal amino acids 1-1357 of the full length dystrophin (Dp427m isoform). The Dp260-2 transcript encodes a unique N-terminal MSARKLRNLSYKK

sequence.

Dystrophin NM 004013.2 26 NP 004004.1 27 Transcript Variant: Dp 140 isoform Dp 140 transcripts use exons 45-79, starting at Sequence Nucleic Acid Nucleic Protein Accession Protein Description

Name Accession No. Acid No. SEQ ID

SEQ ID NO:

NO:

a promoter/exon 1 located in intron 44. Dp 140 transcripts have a long (l kb) 5' UTR since translation is initiated in exon 51 (corresponding to aa 2461 of dystrophin). In addition to the alternative promoter and exon 1, differential splicing of exons 71-74 and 78 produces at least five Dpl40 isoforms. Of these, this transcript (Dp 140) contains all of the exons.

Dystrophin NM 004014.2 28 NP 004005.1 29 Transcript Variant: Dpi 16 isoform transcript Dpi 16 uses exons 56-79, starting from a promoter/exon 1 within intron 55. As a result, the Dp 116 isoform contains a unique N-terminal MLHRKTYHVK aa sequence, instead of aa 1-2739 of dystrophin. Differential splicing produces several Dpl l6-subtypes. The Dpi 16 isoform is also known as S-dystrophin or apo-dystrophin-2.

Dystrophin NM 004015.2 30 NP 004006.1 31 Transcript Variant: Dp71 isoform Dp71 transcripts use exons 63-79 with a novel 80- to 100-nt exon containing an ATG start site for a new coding sequence of 17 nt. The short coding sequence is in-frame with the consecutive dystrophin sequence from exon 63.

Differential splicing of exons 71 and 78 produces at least four Dp71 isoforms. Of these, this transcript (Dp71) includes both exons 71 and 78.

Dystrophin NM 004016.2 32 NP 004007.1 33 Transcript Variant: Dp7 lb isoform Dp71 transcripts use exons 63-79 with a Sequence Nucleic Acid Nucleic Protein Accession Protein Description

Name Accession No. Acid No. SEQ ID

SEQ ID NO:

NO:

novel 80- to 100-nt exon containing an ATG start site for a new coding sequence of 17 nt. The short coding sequence is in-frame with the consecutive dystrophin sequence from exon 63.

Differential splicing of exons 71 and 78 produces at least four Dp71 isoforms. Of these, this transcript (Dp71b) lacks exon 78 and encodes a protein with a different C- terminus than Dp71 and Dp7 la isoforms.

Dystrophin NM 004017.2 34 NP 004008.1 35 Transcript Variant: Dp71a isoform Dp71 transcripts use exons 63-79 with a novel 80- to 100-nt exon containing an ATG start site for a new coding sequence of 17 nt. The short coding sequence is in-frame with the consecutive dystrophin sequence from exon 63.

Differential splicing of exons 71 and 78 produces at least four Dp71 isoforms. Of these, this transcript (Dp71a) lacks exon 71.

Dystrophin NM 004018.2 36 NP 004009.1 37 Transcript Variant: Dp71ab isoform Dp71 transcripts use exons 63-79 with a novel 80- to 100-nt exon containing an ATG start site for a new coding sequence of 17 nt. The short coding sequence is in-frame with the consecutive dystrophin sequence from exon 63.

Differential splicing of exons 71 and 78 produces at least four Dp71 isoforms. Of these, this transcript (Dp71ab) lacks both exons 71 and 78 and Sequence Nucleic Acid Nucleic Protein Accession Protein Description

Name Accession No. Acid No. SEQ ID

SEQ ID NO:

NO:

and 78 produces at least five Dpl40 isoforms. Of these, this transcript (Dp 140b) lacks exon 78 and encodes a protein with a unique C- terminus.

Dystrophin NM 004022.2 44 NP 004013.1 45 Transcript Variant:

Dpl40ab Dp 140 transcripts use isoform exons 45-79, starting at a promoter/exon 1 located in intron 44. Dp 140 transcripts have a long (l kb) 5' UTR since translation is initiated in exon 51 (corresponding to aa 2461 of dystrophin). In addition to the alternative promoter and exon 1, differential splicing of exons 71-74 and 78 produces at least five Dpl40 isoforms. Of these, this transcript (Dpl40ab) lacks exons 71 and 78 and encodes a protein with a unique C -terminus.

Dystrophin NM 004023.2 46 NP 004014.1 47 Transcript Variant:

Dpl40bc Dp 140 transcripts use isoform exons 45-79, starting at a promoter/exon 1 located in intron 44. Dp 140 transcripts have a long (l kb) 5' UTR since translation is initiated in exon 51 (corresponding to aa 2461 of dystrophin). In addition to the alternative promoter and exon 1, differential splicing of exons 71-74 and 78 produces at least five Dpl40 isoforms. Of these, this transcript (Dpl40bc) lacks exons 71-74 and 78 and encodes a protein with a unique C-terminus.

Dystrophin XM 006724469.3 48 XP 006724532.1 49

isoform X2

Dystrophin XM 011545467.1 50 XP 011543769.1 51

isoform X5 Sequence Nucleic Acid Nucleic Protein Accession Protein Description

Name Accession No. Acid No. SEQ ID

SEQ ID NO:

NO:

Dystrophin XM 006724473.2 52 XP 006724536.1 53

isoform X6

Dystrophin XM 006724475.2 54 XP 006724538.1 55

isoform X8

Dystrophin XM 017029328.1 56 XP 016884817.1 57

isoform X4

Dystrophin XM 006724468.2 58 XP 006724531.1 59

isoform XI

Dystrophin XM 017029331.1 60 XP 016884820.1 61

isoform XI 3

Dystrophin XM 006724470.3 62 XP 006724533.1 63

isoform X3

Dystrophin XM 006724474.3 64 XP 006724537.1 65

isoform X7

Dystrophin XM 011545468.2 66 XP 011543770.1 67

isoform X9

Dystrophin XM 017029330.1 68 XP 016884819.1 69

isoform XI 1

Dystrophin XM 017029329.1 70 XP 016884818.1 71

isoform X10

Dystrophin XM 011545469.1 72 XP 011543771.1 73

isoform X12

The murine dystrophin protein has the following amino acid sequence (Uniprot

Accession No. PI 1531, SEQ. ID. NO. 74):

1 MWWVDCYRDV KKTTKWNASK GKHDNSDDGK RDGTGKKKGS TRVHANNV K ARVKNNVDVN

61 GSTDVDGNHK TGWNHWVKNV MKTMAGTNSK SWVRSTRNYV NV TSSWSDG ANAHSHRDDW

121 NSWSHSATR HANAKCGKDD VATTYDKKSM YTSWSAVMR TSSKVTRHHH MHYSTVSAGY

181 TSSSKRKSYA TAAYVATSDS TSYSHARDKS DSSMTV DSY TAVSWSADTR AGSNDWKHA

241 HGMMDTSHGV GNVGSVGKGK SDAVM NSRW CRVASMKSKH KVMDNKKDDW TKTRTKKMGD

301 DKCVHKVDVR V STHMWW DSSGDHATAA KVGDRWANCR WTDRWVDKWH TCSTWSKDAM

361 KNTSGKDNMM SSHKSTKDKK KTMKSSNDSA KNKSVTKMWM NARWDNTKKS SASAVTTTST

421 TTVMTVTMVT TRMVKHAKKR TVDSRKRDVD THSWTRSAVS SAVYRKGNSD KVNAARKAKR

481 KDASRSAAVM ANGVNASRAS NSRWTCSRV WYTNTYNMTT TANKTSTTST AKSKCKDVNR

541 SAKSKKGGMD ADVATNHNHD GVRAKKTDTM RYTMSSRTWS SKSVYSVTYM RGKASSKNGN

601 YSDTVKMAKK ASCKYSGHWK KSSVSCKHM KRKNHKTKWM AVDVKWAGDA KKKCRVGDTS

661 NSVNGGKKSA ASRTRNTWDH CRVYTRKAKA GDKTVSKDSM HWMTAYRDYK TDTAVMKRAK

721 AKTKVKTTVN SVAHASAAKK TTTNYWCTRN GKCKTVWACW HSYKANKWNV KKTMNVAGTV

781 SNMHHSNNRA TTDGGVMDNT NSRWRHAVRK KSSAKSHSDK AAYTDKVDAA MAKSDTSHSM

841 KKHNGKDANR VSDVAKKDVS MKRKANRSKM DVKMHATKSV VSSHCVNYKS SVKSVMVKTG

901 RVKKTNKDRV TAKHYNGAKV TRKKCKSRKM RKMNVTWAAT DTTKRSAVGM SNDSVAWGKA

961 TKKKAHKSVT GSKMVGKKTV DKSNSNWAVT SRVWNYKHMT DNTKWHADDS KKKKDKRKAM

1021 NDMRKVDSTR DAAKMANRGD HCRKWSNRR AASHRKTGKA SKNSDKAGVN KDNKDMSDNG

1081 TV RGDNRTD RKRKKTKHNA KDRSRRKKAS HWYYKRADDK CDKKASRDRK KDRKKKNAVR 1141 RAGSNGAAMA VT SKRWRSNA RRNAHTHTMV VTTDMDVSYV STYTSHASVD HNTCAKDDKS

12 01 KNKDNSGRDH KKKTAASATS MKVKVAVAMD GKHRMYKRGR DRSVKWRHHY DMKVNWNVKK

12 61 TNNWHAKYKW YKDGGRAWR TNATGS SKTD VNKGS SRWHD CKARRKRKNV S RDNVWADNA

1321 TGDKVKARGK NTGGAWSAR DKKKKTNWKV S RAKGVHKDR DHWSRNYNSA GDKVTVHGKA

13 81 DVRSKGHYKK STVKRKDRSW AVNHRRTKDR AGSTTGASAS TVTVT SWTK TVSKMS SVAA

14 41 DNRAWTTDWS DRVKSRVMVG DDNMKKATDR RTAANKNKT S NARTTDRRWD VNRRNMKDST

15 01 WAKAVGVRGK DSWKGHTVDA KKTTKAKDRR SVDVANDAKR DYSADDTRKV HMTNNT SWGN

15 61 HKRVSAATHR DKSWTATTAN VDAS RKKDSR GVRMKWDGTH TDYHNDNGKR S GSDARRDNM

1 621 NKWSKKSNRS HAS S DWKRHS VWKDDS RAGG DAVKNDHRAK RKTKVMSTTV RTGKYRRANV

1 681 TRRKAV AWD KNRSADWRKD ARAADDKRAV KGSWVGDDS D HKVKARGAKN VNRVNDAHTT

17 41 GSYNSTDNTR WRVAVDRVRH AHRDGASHST SVGWRASNKV YYNHTTTCWD HKMTYSADNN

1 8 01 VRSAYRTAMK RRKACDS SAA CDADHNKNDM DNCTTYDRHN NV VCVDMCN WNVYDTGRTG

1 8 61 RRVSKTGSCK AHDKYRYKVA S STGCDRRGH DSRGVAS GGS NSVRS CANNK AADWMR SMVW

1 921 VHRVAAATAK HAKCNCKCGR YRSKHNYDCS CSGRVAKGHK MHYMVYCTTT S GDVRDAKVK

1 9 81 NKRTKRYAKH RMGYVTVGDN MTVTNWVDSA AS S SHDDTHS RHYAS RAM S NGSYNDS SNS

2 041 DDHHYCSNDS SRSAS S RGRA DNRNAYDRKH HKGS SMMTS S RDAAAKRHKG RARMDHNKSH

2 1 01 RRAAKV GTT VS S STS RS DS SMRWGST S S MGDSDTSTGV MNNS S SRGRN AGKMRDTM

Dystrophin is an important component within muscle tissue that provides structural stability to the dystroglycan complex (DGC) of the cell membrane. While both sexes can carry the mutation, females are rarely affected with the skeletal muscle form of the disease.

Mutations vary in nature and frequency. Large genetic deletions are found in about 60- 70% of cases, large duplications are found in about 10% of cases, and point mutants or other small changes account for about 15-30% of cases. One study examined some 7000 mutations and catalogued a total of 5,682 large mutations (80% of total mutations), of which 4,894 (86%) were deletions (1 exon or larger) and 784 (14%) were duplications (1 exon or larger). There were 1,445 small mutations (smaller than 1 exon, 20% of all mutations), of which 358 (25%) were small deletions and 132 (9%) small insertions, while 199 (14%) affected the splice sites. Point mutations totaled 756 (52% of small mutations) with 726 (50%) nonsense mutations and 30 (2%) missense mutations. Finally, 22 (0.3%) mid-intronic mutations were observed. In addition, mutations were identified within the database that would potentially benefit from novel genetic therapies for DMD including stop codon read-through therapies (10% of total mutations) and exon skipping therapy (80% of deletions and 55% of total mutations).

B. Symptoms

Symptoms usually appear in boys between the ages of 2 and 3 and may be visible in early infancy. Even though symptoms do not appear until early infancy, laboratory testing can identify children who carry the active mutation at birth. Progressive proximal muscle weakness of the legs and pelvis associated with loss of muscle mass is observed first. Eventually this weakness spreads to the arms, neck, and other areas. Early signs may include pseudohypertrophy (enlargement of calf and deltoid muscles), low endurance, and difficulties in standing unaided or inability to ascend staircases. As the condition progresses, muscle tissue experiences wasting and is eventually replaced by fat and fibrotic tissue (fibrosis). By age 10, braces may be required to aid in walking but most patients are wheelchair dependent by age 12. Later symptoms may include abnormal bone development that lead to skeletal deformities, including curvature of the spine. Due to progressive deterioration of muscle, loss of movement occurs, eventually leading to paralysis. Intellectual impairment may or may not be present but if present, does not progressively worsen as the child ages. The average life expectancy for males afflicted with DMD is around 25.

The main symptom of Duchenne muscular dystrophy, a progressive neuromuscular disorder, is muscle weakness associated with muscle wasting with the voluntary muscles being first affected, especially those of the hips, pelvic area, thighs, shoulders, and calves. Muscle weakness also occurs later, in the arms, neck, and other areas. Calves are often enlarged. Symptoms usually appear before age 6 and may appear in early infancy. Other physical symptoms are:

• Awkward manner of walking, stepping, or running - (patients tend to walk on their forefeet, because of an increased calf muscle tone. Also, toe walking is a compensatory adaptation to knee extensor weakness.)

• Frequent falls

• Fatigue

• Difficulty with motor skills (running, hopping, jumping)

• Lumbar hyperlordosis, possibly leading to shortening of the hip-flexor muscles. This has an effect on overall posture and a manner of walking, stepping, or running.

• Muscle contractures of Achilles tendon and hamstrings impair functionality because the muscle fibers shorten and fibrose in connective tissue

• Progressive difficulty walking

• Muscle fiber deformities

• Pseudohypertrophy (enlarging) of tongue and calf muscles. The muscle tissue is eventually replaced by fat and connective tissue, hence the term pseudohypertrophy. • Higher risk of neurobehavioral disorders (e.g. , ADHD), learning disorders (dyslexia), and non-progressive weaknesses in specific cognitive skills (in particular short-term verbal memory), which are believed to be the result of absent or dysfunctional dystrophin in the brain.

Eventual loss of ability to walk (usually by the age of 12)

• Skeletal deformities (including scoliosis in some cases)

• Trouble getting up from lying or sitting position

The condition can often be observed clinically from the moment the patient takes his first steps, and the ability to walk usually completely disintegrates between the time the patient is 9 to 12 years of age. Most men affected with DMD become essentially "paralyzed from the neck down" by the age of 21. Muscle wasting begins in the legs and pelvis, then progresses to the muscles of the shoulders and neck, followed by loss of arm muscles and respiratory muscles. Calf muscle enlargement (pseudohypertrophy) is quite obvious. Cardiomyopathy particularly (dilated cardiomyopathy) is common, but the development of congestive heart failure or arrhythmia (irregular heartbeat) is only occasional.

A positive Gowers' sign reflects the more severe impairment of the lower extremities muscles. The child helps himself to get up with upper extremities: first by rising to stand on his arms and knees, and then "walking" his hands up his legs to stand upright. Affected children usually tire more easily and have less overall strength than their peers. Creatine kinase (CPK- MM) levels in the bloodstream are extremely high. An electromyography (EMG) shows that weakness is caused by destruction of muscle tissue rather than by damage to nerves. Genetic testing can reveal genetic errors in the Xp21 gene. A muscle biopsy (immunohistochemistry or immunoblotting) or genetic test (blood test) confirms the absence of dystrophin, although improvements in genetic testing often make this unnecessary.

Other symptoms include:

• Abnormal heart muscle (cardiomyopathy)

• Congestive heart failure or irregular heart rhythm (arrhythmia)

• Deformities of the chest and back (scoliosis)

• Enlarged muscles of the calves, buttocks, and shoulders (around age 4 or 5). These muscles are eventually replaced by fat and connective tissue (pseudohypertrophy).

• Loss of muscle mass (atrophy)

• Muscle contractures in the heels, legs • Muscle deformities

• Respiratory disorders, including pneumonia and swallowing with food or fluid passing into the lungs (in late stages of the disease)

C. Causes

Duchenne muscular dystrophy (DMD) is caused by a mutation of the dystrophin gene at locus Xp21, located on the short arm of the X chromosome. Dystrophin is responsible for connecting the cytoskeleton of each muscle fiber to the underlying basal lamina (extracellular matrix), through a protein complex containing many subunits. The absence of dystrophin permits excess calcium to penetrate the sarcolemma (the cell membrane). Alterations in calcium and signaling pathways cause water to enter into the mitochondria, which then burst.

In skeletal muscle dystrophy, mitochondrial dysfunction gives rise to an amplification of stress-induced cytosolic calcium signals and an amplification of stress-induced reactive- oxygen species (ROS) production. In a complex cascading process that involves several pathways and is not clearly understood, increased oxidative stress within the cell damages the sarcolemma and eventually results in the death of the cell. Muscle fibers undergo necrosis and are ultimately replaced with adipose and connective tissue.

DMD is inherited in an X-linked recessive pattern. Females will typically be carriers for the disease while males will be affected. Typically, a female carrier will be unaware they carry a mutation until they have an affected son. The son of a carrier mother has a 50% chance of inheriting the defective gene from his mother. The daughter of a carrier mother has a 50% chance of being a carrier and a 50% chance of having two normal copies of the gene. In all cases, an unaffected father will either pass a normal Y to his son or a normal X to his daughter. Female carriers of an X-linked recessive condition, such as DMD, can show symptoms depending on their pattern of X-inactivation.

Exon deletions preceding exon 51 of the human DMD gene, which disrupt the open reading frame (ORF) by juxtaposing out of frame exons, represent the most common type of human DMD mutation. Skipping of exon 51 can, in principle, restore the DMD ORF in 13% of DMD patients with exon deletions.

Duchenne muscular dystrophy has an incidence of 1 in 5000 male infants. Mutations within the dystrophin gene can either be inherited or occur spontaneously during germline transmission. An exemplary but non-limiting mutation is deletion of exons 8 and 9 and a corresponding iPSC model is ΔΕχ8-9 iDMD and a mouse model is ΔΕχ8-9 DMD. D. Diagnosis

Genetic counseling is advised for people with a family history of the disorder.

Duchenne muscular dystrophy can be detected with about 95% accuracy by genetic studies performed during pregnancy.

DNA test. The muscle-specific isoform of the dystrophin gene is composed of 79 exons, and DNA testing and analysis can usually identify the specific type of mutation of the exon or exons that are affected. DNA testing confirms the diagnosis in most cases.

Muscle biopsy. If DNA testing fails to find the mutation, a muscle biopsy test may be performed. A small sample of muscle tissue is extracted (usually with a scalpel instead of a needle) and a dye is applied that reveals the presence of dystrophin. Complete absence of the protein indicates the condition.

Over the past several years DNA tests have been developed that detect more of the many mutations that cause the condition, and muscle biopsy is not required as often to confirm the presence of Duchenne's.

Prenatal tests. DMD is carried by an X-linked recessive gene. Males have only one X chromosome, so one copy of the mutated gene will cause DMD. Fathers cannot pass X-linked traits on to their sons, so the mutation is transmitted by the mother.

If the mother is a carrier, and therefore one of her two X chromosomes has a DMD mutation, there is a 50% chance that a female child will inherit that mutation as one of her two X chromosomes, and be a carrier. There is a 50% chance that a male child will inherit that mutation as his one X chromosome, and therefore have DMD.

Prenatal tests can tell whether an unborn child has the most common mutations. There are many mutations responsible for DMD, and some have not been identified, so genetic testing only works when family members with DMD have a mutation that has been identified.

Prior to invasive testing, determination of the fetal sex is important; while males are sometimes affected by this X-linked disease, female DMD is extremely rare. This can be achieved by ultrasound scan at 16 weeks or more recently by free fetal DNA testing. Chorion villus sampling (CVS) can be done at 11-14 weeks, and has a 1% risk of miscarriage.

Amniocentesis can be done after 15 weeks, and has a 0.5% risk of miscarriage. Fetal blood sampling can be done at about 18 weeks. Another option in the case of unclear genetic test results is fetal muscle biopsy. E. Treatment

There is no current cure for DMD, and an ongoing medical need has been recognized by regulatory authorities. Phase l-2a trials with exon skipping treatment for certain mutations have halted decline and produced small clinical improvements in walking. Treatment is generally aimed at controlling the onset of symptoms to maximize the quality of life, and include the following:

• Corticosteroids such as prednisolone and deflazacort increase energy and strength and defer severity of some symptoms.

• Randomized control trials have shown that beta-2-agonists increase muscle strength but do not modify disease progression. Follow-up time for most RCTs on beta2-agonists is only around 12 months and hence results cannot be extrapolated beyond that time frame.

• Mild, non-jarring physical activity such as swimming is encouraged. Inactivity (such as bed rest) can worsen the muscle disease.

• Physical therapy is helpful to maintain muscle strength, flexibility, and function.

• Orthopedic appliances (such as braces and wheelchairs) may improve mobility and the ability for self-care. Form-fitting removable leg braces that hold the ankle in place during sleep can defer the onset of contractures.

• Appropriate respiratory support as the disease progresses is important.

Comprehensive multi-disciplinary care standards/guidelines for DMD have been developed by the Centers for Disease Control and Prevention (CDC), and were published in two parts and are available at world-wide-web at treat-nmd.eu/dmd/care/diagnosis-management-DMD.

DMD generally progresses through five stages, as outlined in Bushby et al, Lancet Neurol, 9(1): 77-93 (2010) and Bushby et al, Lancet Neurol, 9(2): 177-198 (2010), incorporated by reference in their entireties. During the presymptomatic stage, patients typically show developmental delay, but no gait disturbance. During the early ambulatory stage, patients typically show the Gowers' sign, waddling gait, and toe walking. During the late ambulatory stage, patients typically exhibit an increasingly labored gait and begin to lose the ability to climb stairs and rise from the floor. During the early non-ambulatory stage, patients are typically able to self-propel for some time, are able to maintain posture, and may develop scoliosis. During the late non-ambulatory stage, upper limb function and postural maintenance is increasingly limited. In some embodiments, treatment is initiated in the presymptomatic stage of the disease. In some embodiments, treatment is initiated in the early ambulatory stage. In some embodiments, treatment is initiated in the late ambulatory stage. In embodiments, treatment is initiated during the early non-ambulatory stage. In embodiments, treatment is initiated during the late non-ambulatory stage.

1. Physical Therapy

Physical therapists are concerned with enabling patients to reach their maximum physical potential. Their aim is to:

• minimize the development of contractures and deformity by developing a program of stretches and exercises where appropriate

• anticipate and minimize other secondary complications of a physical nature by recommending bracing and durable medical equipment

• monitor respiratory function and advise on techniques to assist with breathing exercises and methods of clearing secretions

2. Respiration Assistance

Modern "volume ventilators/respirators," which deliver an adjustable volume (amount) of air to the person with each breath, are valuable in the treatment of people with muscular dystrophy related respiratory problems. The ventilator may require an invasive endotracheal or tracheotomy tube through which air is directly delivered, but, for some people non-invasive delivery through a face mask or mouthpiece is sufficient. Positive airway pressure machines, particularly bi-level ones, are sometimes used in this latter way. The respiratory equipment may easily fit on a ventilator tray on the bottom or back of a power wheelchair with an external battery for portability.

Ventilator treatment may start in the mid to late teens when the respiratory muscles can begin to collapse. If the vital capacity has dropped below 40% of normal, a volume ventilator/respirator may be used during sleeping hours, a time when the person is most likely to be under ventilating ("hypoventilating"). Hypoventilation during sleep is determined by a thorough history of sleep disorder with an oximetry study and a capillary blood gas. A cough assist device can help with excess mucus in lungs by hyperinflation of the lungs with positive air pressure, then negative pressure to get the mucus up. If the vital capacity continues to decline to less than 30 percent of normal, a volume ventilator/respirator may also be needed during the day for more assistance. The person gradually will increase the amount of time using the ventilator/respirator during the day as needed.

F. Prognosis

Duchenne muscular dystrophy is a progressive disease which eventually affects all voluntary muscles and involves the heart and breathing muscles in later stages. The life expectancy is currently estimated to be around 25, but this varies from patient to patient. Recent advancements in medicine are extending the lives of those afflicted. The Muscular Dystrophy Campaign, which is a leading UK charity focusing on all muscle disease, states that "with high standards of medical care young men with Duchenne muscular dystrophy are often living well into their 30s."

In rare cases, persons with DMD have been seen to survive into the forties or early fifties, with the use of proper positioning in wheelchairs and beds, ventilator support (via tracheostomy or mouthpiece), airway clearance, and heart medications, if required. Early planning of the required supports for later-life care has shown greater longevity in people living with DMD.

Curiously, in the mdx mouse model of Duchenne muscular dystrophy, the lack of dystrophin is associated with increased calcium levels and skeletal muscle myonecrosis. The intrinsic laryngeal muscles (ILM) are protected and do not undergo myonecrosis. ILM have a calcium regulation system profile suggestive of a better ability to handle calcium changes in comparison to other muscles, and this may provide a mechanistic insight for their unique pathophysiological properties. The ILM may facilitate the development of novel strategies for the prevention and treatment of muscle wasting in a variety of clinical scenarios. II. CRISPR Systems

A. CRISPRs

CRISPRs (clustered regularly interspaced short palindromic repeats) are DNA loci containing short repetitions of base sequences. Each repetition is followed by short segments of "spacer DNA" from previous exposures to a virus. CRISPRs are found in approximately 40% of sequenced eubacteria genomes and 90% of sequenced archaea. CRISPRs are often associated with Cas genes that code for proteins related to CRISPRs. The CRISPR/Cas system is a prokaryotic immune system that confers resistance to foreign genetic elements such as plasmids and phages and provides a form of acquired immunity. CRISPR spacers recognize and silence these exogenous genetic elements like RNAi in eukaryotic organisms. CRISPR repeats range in size from 24 to 48 base pairs. They usually show some dyad symmetry, implying the formation of a secondary structure such as a hairpin, but are not truly palindromic. Repeats are separated by spacers of similar length. Some CRISPR spacer sequences exactly match sequences from plasmids and phages, although some spacers match the prokary ote' s genome (self-targeting spacers). New spacers can be added rapidly in response to phage infection.

B. Cas Nucleases

CRISPR-associated (cas) genes are often associated with CRISPR repeat-spacer arrays. As of 2013, more than forty different Cas protein families had been described. Of these protein families, Casl appears to be ubiquitous among different CRISPR/Cas systems. Particular combinations of cas genes and repeat structures have been used to define 8 CRISPR subtypes (Ecoli, Ypest, Nmeni, Dvulg, Tneap, Hmari, Apem, and Mtube), some of which are associated with an additional gene module encoding repeat-associated mysterious proteins (RAMPs). More than one CRISPR subtype may occur in a single genome. The sporadic distribution of the CRISPR/Cas subtypes suggests that the system is subject to horizontal gene transfer during microbial evolution.

Exogenous DNA is apparently processed by proteins encoded by Cas genes into small elements (-30 base pairs in length), which are then somehow inserted into the CRISPR locus near the leader sequence. RNAs from the CRISPR loci are constitutively expressed and are processed by Cas proteins to small RNAs composed of individual, exogenously-derived sequence elements with a flanking repeat sequence. The RNAs guide other Cas proteins to silence exogenous genetic elements at the RNA or DNA level. Evidence suggests functional diversity among CRISPR subtypes. The Cse (Cas subtype Ecoli) proteins (called CasA-E in E. coli) form a functional complex, Cascade, that processes CRISPR RNA transcripts into spacer- repeat units that Cascade retains. In other prokary otes, Cas6 processes the CRISPR transcripts. Interestingly, CRISPR-based phage inactivation in E. coli requires Cascade and Cas3, but not Casl and Cas2. The Cmr (Cas RAMP module) proteins found in Pyrococcus furiosus and other prokary otes form a functional complex with small CRISPR RNAs that recognizes and cleaves complementary target RNAs. RNA-guided CRISPR enzymes are classified as type V restriction enzymes.

Cas9 is a nuclease, an enzyme specialized for cutting DNA, with two active cutting sites, one for each strand of the double helix. The team demonstrated that they could disable one or both sites while preserving Cas9's ability to locate its target DNA. TracrRNA and spacer RNA can be combined into a "single-guide RNA" molecule that, mixed with Cas9, can find and cut the correct DNA targets, and such synthetic guide RNAs are able to be used for gene editing.

Cas9 proteins are highly enriched in pathogenic and commensal bacteria. CRISPR/Cas- mediated gene regulation may contribute to the regulation of endogenous bacterial genes, particularly during bacterial interaction with eukaryotic hosts. For example, Cas protein Cas9 of Francisella novicida uses a unique, small, CRISPR/Cas-associated RNA (scaRNA) to repress an endogenous transcript encoding a bacterial lipoprotein that is critical for F. novicida to dampen host response and promote virulence. One group showed that coinjection of Cas9 mRNA and sgRNAs into the germline (zygotes) generated nice with mutations. Delivery of Cas9 DNA sequences also is contemplated.

The systems CRISPR/Cas are separated into three classes. Class 1 uses several Cas proteins together with the CRISPR RNAs (crRNA) to build a functional endonuclease. Class 2 CRISPR systems use a single Cas protein with a crRNA. Cpfl has been recently identified as a Class II, Type V CRISPR/Cas systems containing a 1,300 amino acid protein. See also U.S. Patent Publication 2014/0068797, which is incorporated by reference in its entirety.

In some embodiments, the compositions of the disclosure include a small version of a Cas9 from the bacterium Staphylococcus aureus (UniProt Accession No. J7RUA5). The small version of the Cas9 provides advantages over wild type or full length Cas9. In some embodiments the Cas9 is a spCas9 (AddGene).

C. Cpfl Nucleases

Clustered Regularly Interspaced Short Palindromic Repeats from Prevotella and Francisella 1 or CRISPR/Cpfl is a DNA-editing technology which shares some similarities with the CRISPR/Cas9 system. Cpfl is an RNA-guided endonuclease of a class II CRISPR/Cas system. This acquired immune mechanism is found in Prevotella and Francisella bacteria. It prevents genetic damage from viruses. Cpfl genes are associated with the CRISPR locus, coding for an endonuclease that use a guide RNA to find and cleave viral DNA. Cpfl is a smaller and simpler endonuclease than Cas9, overcoming some of the CRISPR/Cas9 system limitations.

Cpfl appears in many bacterial species. The ultimate Cpfl endonuclease that was developed into a tool for genome editing was taken from one of the first 16 species known to harbor it. In embodiments, the Cpfl is a Cpfl enzyme from Acidaminococcus (species BV3L6, UniProt Accession No. U2UMQ6; SEQ ID NO. 75), having the sequence set forth below:

1 MTQFEGFTNL YQVSKTLRFE LIPQGKTLKH IQEQGFIEED KARNDHYKEL KPIIDRIYKT

61 YADQCLQLVQ LDWENLSAAI DSYRKEKTEE TRNALIEEQA TYRNAIHDYF IGRTDNLTDA

121 INKRHAEIYK GLFKAELFNG KVLKQLGTVT TTEHENALLR SFDKFTTYFS GFYENRKNVF

181 SAEDISTAIP HRIVQDNFPK FKENCHI FTR LITAVPSLRE HFENVKKAIG IFVSTSIEEV

241 FSFPFYNQLL TQTQIDLYNQ LLGGI SREAG TEKIKGLNEV LNLAIQKNDE TAHIIASLPH

301 RFIPLFKQIL SDRNTLSFIL EEFKSDEEVI QSFCKYKTLL RNENVLETAE ALFNELNSID

361 LTHIFISHKK LETISSALCD HWDTLRNALY ERRI SELTGK ITKSAKEKVQ RSLKHEDINL

421 QEIISAAGKE LSEAFKQKTS EILSHAHAAL DQPLPTTLKK QEEKEILKSQ LDSLLGLYHL

481 LDWFAVDESN EVDPEFSARL TGIKLEMEPS LSFYNKARNY ATKKPYSVEK FKLNFQMPTL

541 ASGWDVNKEK NNGAILFVKN GLYYLGIMPK QKGRYKALSF EPTEKTSEGF DKMYYDYFPD

601 AAKMIPKCST QLKAVTAHFQ THTTPILLSN NFIEPLEITK EIYDLNNPEK EPKKFQTAYA

661 KKTGDQKGYR EALCKWIDFT RDFLSKYTKT TSIDLSSLRP SSQYKDLGEY YAELNPLLYH

721 ISFQRIAEKE IMDAVETGKL YLFQIYNKDF AKGHHGKPNL HTLYWTGLFS PENLAKTSIK

781 LNGQAELFYR PKSRMKRMAH RLGEKMLNKK LKDQKTPI PD TLYQELYDYV NHRLSHDLSD

841 EARALLPNVI TKEVSHEIIK DRRFTSDKFF FHVPITLNYQ AANSPSKFNQ RVNAYLKEHP

901 ETPI IGIDRG ERNLIYITVI DSTGKILEQR SLNTIQQFDY QKKLDNREKE RVAARQAWSV

961 VGTIKDLKQG YLSQVIHEIV DLMIHYQAW VLENLNFGFK SKRTGIAEKA VYQQFEKMLI

1021 DKLNCLVLKD YPAEKVGGVL NPYQLTDQFT SFAKMGTQSG FLFYVPAPYT SKIDPLTGFV

1081 DPFVWKTIKN HESRKHFLEG FDFLHYDVKT GDFILHFKMN RNLSFQRGLP GFMPAWDIVF

1141 EKNETQFDAK GTPFIAGKRI VPVIENHRFT GRYRDLYPAN ELIALLEEKG IVFRDGSNIL

1201 PKLLENDDSH AIDTMVALIR SVLQMRNSNA ATGEDYINSP VRDLNGVCFD SRFQNPEWPM

1261 DADANGAYHI ALKGQLLLNH LKESKDLKLQ NGISNQDWLA YIQELRN

In some embodiments, the Cpfl is a Cpfl enzyme from Lachnospiraceae (species ND2006, UniProt Accession No. A0A182DWE3; SEQ ID NO. 76), having the sequence set forth below:

1 AASKLEKFTN CYSLSKTLRF KAIPVGKTQE NIDNKRLLVE DEKRAEDYKG VKKLLDRYYL 61 SFINDVLHSI KLKNLNNYIS LFRKKTRTEK ENKELENLEI NLRKEIAKAF KGAAGYKSLF 121 KKDIIETILP EAADDKDEIA LVNSFNGFTT AFTGFFDNRE NMFSEEAKST SIAFRCINEN 181 LTRYI SNMDI FEKVDAIFDK HEVQEIKEKI LNSDYDVEDF FEGEFFNFVL TQEGIDVYNA 241 I IGGFVTESG EKIKGLNEYI NLYNAKTKQA LPKFKPLYKQ VLSDRESLSF YGEGYTSDEE 301 VLEVFRNTLN KNSEIFSSIK KLEKLFKNFD EYSSAGIFVK NGPAISTISK DIFGEWNLIR 361 DKWNAEYDDI HLKKKAWTE KYEDDRRKSF KKIGSFSLEQ LQEYADADLS WEKLKEIII 421 QKVDEIYKVY GSSEKLFDAD FVLEKSLKKN DAWAIMKDL LDSVKSFENY IKAFFGEGKE

481 TNRDESFYGD FVLAYDILLK VDHIYDAIRN YVTQKPYSKD KFKLYFQNPQ FMGGWDKDKE

541 TDYRATILRY GSKYYLAIMD KKYAKCLQKI DKDDVNGNYE KINYKLLPGP NKMLPKVFFS

601 KKWMAYYNPS EDIQKIYKNG TFKKGDMFNL NDCHKLIDFF KDSISRYPKW SNAYDFNFSE

661 TEKYKDIAGF YREVEEQGYK VSFESASKKE VDKLVEEGKL YMFQIYNKDF SDKSHGTPNL

721 HTMYFKLLFD ENNHGQIRLS GGAELFMRRA SLKKEELWH PANSPIANKN PDNPKKTTTL

781 SYDVYKDKRF SEDQYELHIP IAINKCPKNI FKINTEVRVL LKHDDNPYVI GIDRGERNLL

841 YIVWDGKGN IVEQYSLNEI INNFNGIRIK TDYHSLLDKK EKERFEARQN WTSIENIKEL

901 KAGYI SQWH KICELVEKYD AVIALEDLNS GFKNSRVKVE KQVYQKFEKM LIDKLNYMVD

961 KKSNPCATGG ALKGYQITNK FESFKSMSTQ NGFIFYIPAW LTSKIDPSTG FVNLLKTKYT

1021 SIADSKKFIS SFDRIMYVPE EDLFEFALDY KNFSRTDADY IKKWKLYSYG NRIRIFAAAK

1081 KNNVFAWEEV CLTSAYKELF NKYGINYQQG DIRALLCEQS DKAFYSSFMA LMSLMLQMRN

1141 SITGRTDVDF LISPVKNSDG IFYDSRNYEA QENAILPKNA DANGAYNIAR KVLWAIGQFK

1201 KAEDEKLDKV KIAI SNKEWL EYAQTSVK

In some embodiments, the Cpfl is codon optimized for expression in mammalian cells. In some embodiments, the Cpfl is codon optimized for expression in human cells or mouse cells.

The Cpfl locus contains a mixed alpha/beta domain, a RuvC-I followed by a helical region, a RuvC-II and a zinc finger-like domain. The Cpfl protein has a RuvC-like endonuclease domain that is similar to the RuvC domain of Cas9. Furthermore, Cpfl does not have a HNH endonuclease domain, and the N-terminal of Cpfl does not have the alpha-helical recognition lobe of Cas9.

Cpfl CRISPR-Cas domain architecture shows that Cpfl is functionally unique, being classified as Class 2, type V CRISPR system. The Cpfl loci encode Casl, Cas2 and Cas4 proteins more similar to types I and III than from type II systems. Database searches suggest the abundance of Cpfl -family proteins in many bacterial species.

Functional Cpfl doesn't does not require a tracrRNA. Therefore, functional Cpfl gRNAs of the disclosure may comprise or consist of a crRNA. This benefits genome editing because Cpfl is not only a smaller than Cas9, but also it has a smaller sgRNA molecule (approximately half as many nucleotides as Cas9).

The Cpfl-gRNA (e.g., Cpfl-crRNA) complex cleaves target DNA or RNA by identification of a protospacer adjacent motif 5'-YTN-3' (where "Y" is a pyrimidine and "N" is any nucleobase) or 5'-TTN-3', in contrast to the G-rich PAM targeted by Cas9. After identification of PAM, Cpfl introduces a sticky-end-like DNA double- stranded break of 4 or 5 nucleotides overhang. The CRISPR/Cpfl system comprises or consists of a Cpfl enzyme and a guide RNA that finds and positions the complex at the correct spot on the double helix to cleave target DNA. In its native bacterial hosts, CRISPR/Cpfl systems activity has three stages:

Adaptation, during which Casl and Cas2 proteins facilitate the adaptation of small fragments of DNA into the CRISPR array;

Formation of crRNAs: processing of pre-cr-RNAs producing of mature crRNAs to guide the Cas protein; and

Interference, in which the Cpfl is bound to a crRNA to form a binary complex to identify and cleave a target DNA sequence.

This system has been modified to utilize non-naturally occurring crRNAs, which guide Cpfl to a desired target sequence in a non-bacterial cell, such as a mammalian cell.

B. gRNA

As an RNA guided protein, Cas9 requires a short RNA to direct the recognition of DNA targets. Though Cas9 preferentially interrogates DNA sequences containing a PAM sequence NGG it can bind here without a protospacer target. However, the Cas9-gRNA complex requires a close match to the gRNA to create a double strand break. CRISPR sequences in bacteria are expressed in multiple RNAs and then processed to create guide strands for RNA. Because Eukaryotic systems lack some of the proteins required to process CRISPR RNAs the synthetic construct gRNA was created to combine the essential pieces of RNA for Cas9 targeting into a single RNA expressed with the RNA polymerase type III promoter U6. Synthetic gRNAs are slightly over lOObp at the minimum length and contain a portion which targets the 20 protospacer nucleotides immediately preceding the PAM sequence NGG; gRNAs do not contain a PAM sequence.

In some embodiments, a gRNA targets a site within a wildtype dystrophin gene. In some embodiments, a gRNA targets a site within a mutant dystrophin gene. In some embodiments, a gRNA targets a dystrophin intron. In some embodiments, a gRNA targets a dystrophin exon. In some embodiments, a gRNA targets a site in a dystrophin exon that is expressed and is present in one or more of the dystrophin isoforms shown in Table 3. In some embodiments, a gRNA targets a site in a dystrophin exon that is within the ABD-1 domain of dystrophin. In embodiments, a gRNA targets a dystrophin splice site. In some embodiments, a gRNA targets a splice donor site on the dystrophin gene. In embodiments, a gRNA targets a splice acceptor site on the dystrophin gene. In embodiments, more than one guide RNAs are used to edit a dystrophin gene. In some embodiments, 2, 3, 4, 5, 6, 7, 8, 9, or 10 guide RNAs are used to edit a dystrophin gene. In particular embodiments, two guide RNAs are used to edit a dystrophin gene.

In some embodiments, a first guide RNA targets a first genomic target sequence, and a second guide RNA targets a second genomic target sequence. In some embodiments, the first genomic target sequence and the second genomic target sequence may both be in an intronic region of the dsyrophin gene. In other embodiments, the first genomic target sequence and the second genomic target sequence may both be in an exonic region of the dsyrophin gene. In some embodiments, the first genomic target sequence is an intronic region of the dystropin gene, and the second genomic target sequence is in an exonic region of the dystrophin gene. In some embodiments, the genomic target sequence may be within intron 2, 3, 4, 5, 6, 7, 8, 9, 10, or 1 1 of dystrophin. In some embodiments, the genomic target sequence may be within exon 2, 3, 4, 5, 6, 7, 8, 9, 10, or 11 of dystrophin.

One or more guide RNAs may be used to edit the Actin Binding Domain- 1 region of the dystrophin protein, encoded by exons 2-8 (amino acids 14-240 of SEQ ID NO: 74). For example, a guide RNA may target any one of exons 2-8, such as exon 2, 3, 4, 5, 6, 7 or 8. In some embodiments, a guide RNA may target any one of introns 2-9, such as intron 2, 3, 4, 5, 6, 7, 8, or 9. In some embodiments, a first guide RNA targets any one of exons 2-8, and a second guide RNA targets any one of exons 2-8. In some embodiments, a first guide RNA targets any one of introns 2-9, and a second guide RNA targets any one of introns 2-9. In some embodiments, a first guide RNA targets any one of introns 2-9, and a second guide RNA targets any one of exons 2-8. In some embodiments, a first guide RNA and a second guide RNA target the introns shown in Table 4, or the exons shown in Table 5, below. In these tables, an "x" indicates that the combination shown is contemplated by the instant disclosure.

Table 4: Combinations of Introns Targeted by First and Second gRNA

Table 5: Combinations of Exons Targeted by First and Second gRNA

In some embodiments, a first gRNA targets intron 2 and a second gRNA targets intron 7. In some embodiments, a first gRNA targets intron 5 and a second gRNA targets intron 7. In some embodiments, a first gRNA targets intron 2 and a second gRNA targets intron 9. In some embodiments, a first gRNA targets intron 7 and a second gRNA targets intron 9.

In some embodiments, a first genomic target sequence is located within intron 2 and a second genomic target sequence is located within intron 7. In some embodiments, a first genomic target sequence is located within intron 5 and a second genomic target sequence is located within intron 7. In some embodiments, a first genomic target sequence is located within intron 2 and a second genomic target sequence is located within intron 9. In some embodiments, a first genomic target sequence is located within intron 7 and a second genomic target sequence is located within intron 9.

In some embodiments, one or more gRNAs are used to delete one or more of exons 2-

8. For example, a first gRNA may be targeted to a sequence in an intron 5' to the one or more exons, and a second gRNA may be targeted to a sequence in an intron 3' to the one or more exons. In the presence of a nuclease (e.g. , Cas9 nuclease), the one or more gRNAs cause excision of the one or more exons. In some embodiments, exons 3-7 are deleted. In some embodiments, exons 6-7 are deleted.

Suitable gRNAs for use in various compositions and methods disclosed herein are provided as SEQ ID NOs. 6 to 10. (Table 1). In preferred embodiments, the gRNA is selected from any one of SEQ ID NO. 6 to SEQ ID NO. 10.

In some embodiments, gRNAs of the disclosure comprise a sequence that is complementary to a target sequence within a coding sequence or a non-coding sequence corresponding to the DMD gene, and, therefore, hybridize to the target sequence.

In some embodiments, gRNAs for Cpfl comprise a single crRNA containing a direct repeat scaffold sequence followed by 24 nucleotides of guide sequence. In some embodiments, a "guide" sequence of the crRNA comprises a sequence of the gRNA that is complementary to a target sequence. In some embodiments, crRNA of the disclosure comprises a sequence of the gRNA that is not complementary to a target sequence. "Scaffold" sequences of the disclosure link the gRNA to the Cpfl polypeptide. "Scaffold" sequences of the disclosure are not equivalent to a tracrRNA sequence of a gRNA-Cas9 construct. E. Cas9 versus Cpfl

Cas9 requires two RNA molecules to cut DNA while Cpfl needs one. The proteins also cut DNA at different places, offering researchers more options when selecting an editing site.

Cas9 cuts both strands in a DNA molecule at the same position, leaving behind 'blunt' ends.

Cpfl leaves one strand longer than the other, creating 'sticky' ends that are easier to work with. Cpfl appears to be more able to insert new sequences at the cut site, compared to Cas9.

Although the CRISPR/Cas9 system can efficiently disable genes, it is challenging to insert genes or generate a knock-in. Cpfl lacks tracrRNA, utilizes a T-rich PAM and cleaves DNA via a staggered DNA DSB. In summary, important differences between Cpfl and Cas9 systems are that Cpfl recognizes different P AMs, enabling new targeting possibilities, creates 4-5 nt long sticky ends, instead of blunt ends produced by Cas9, enhancing the efficiency of genetic insertions and specificity during NHEJ or HDR, and cuts target DNA further away from PAM, further away from the Cas9 cutting site, enabling new possibilities for cleaving the DNA.

F. CRISPR/Cpfl-mediated gene editing

The first step in editing the DMD gene using CRISPR/Cpfl is to identify the genomic target sequence. The genomic target for the gRNAs of the disclosure can be any -24 nucleotide DNA sequence within the dystrophin gene, provided that the sequence is unique compared to the rest of the genome.

The next step in editing the DMD gene using CRISPR/Cpfl is to identify all Protospacer Adjacent Motif (PAM) sequences within the genetic region to be targeted. Cpfl utilizes a T-rich PAM sequence (TTTN, wherein N is any nucleotide). The target sequence must be immediately upstream of a PAM. Once all possible PAM sequences and putative target sites have been identified, the next step is to choose which site is likely to result in the most efficient on-target cleavage. The gRNA targeting sequence needs to match the target sequence, and the gRNA targeting sequence must not match additional sites within the genome. In preferred embodiments, the gRNA targeting sequence has perfect homology to the target with no homology elsewhere in the genome. In some embodiments, a given gRNA targeting sequence will have additional sites throughout the genome where partial homology exists. These sites are called "off-targets" and should be considered when designing a gRNA. In general, off-target sites are not cleaved as efficiently when mismatches occur near the PAM sequence, so gRNAs with no homology or those with mismatches close to the PAM sequence will have the highest specificity. In addition to "off-target activity", factors that maximize cleavage of the desired target sequence ("on-target activity") must be considered. It is known to those of skill in the art that two gRNA targeting sequences, each having 100% homology to the target DNA may not result in equivalent cleavage efficiency. In fact, cleavage efficiency may increase or decrease depending upon the specific nucleotides within the selected target sequence. Close examination of predicted on-target and off-target activity of each potential gRNA targeting sequence is necessary to design the best gRNA. Several gRNA design programs have been developed that are capable of locating potential PAM and target sequences and ranking the associated gRNAs based on their predicted on-target and off-target activity (e.g. CRISPRdirect, available at www.crispr.dbcls.jp).

The next step is to synthesize and clone desired gRNAs. Targeting oligos can be synthesized, annealed, and inserted into plasmids containing the gRNA scaffold using standard restriction-ligation cloning. However, the exact cloning strategy will depend on the gRNA vector that is chosen. The gRNAs for Cpfl are notably simpler than the gRNAs for Cas9, and only consist of a single crRNA containing direct repeat scaffold sequence followed by -24 nucleotides of guide sequence. Cpfl requires a minimum of 16 nucleotides of guide sequence to achieve detectable DNA cleavage, and a minimum of 18 nucleotides of guide sequence to achieve efficient DNA cleavage in vitro. In some embodiments, 20-24 nucleotides of guide sequence is used. The seed region of the Cpfl gRNA is generally within the first 5 nucleotides on the 5 ' end of the guide sequence. Cpfl makes a staggered cut in the target genomic DNA. In AsCpfl and LbCpfl, the cut occurs 19 bp after the PAM on the targeted (+) strand, and 23 bp on the other strand.

Each gRNA should then be validated in one or more target cell lines. For example, after the CRISPR and gRNA are delivered to the cell, the genomic target region may be amplified using PCR and sequenced according to methods known to those of skill in the art.

In some embodiments, gene editing may be performed in vitro or ex vivo. In some embodiments, cells are contacted in vitro or ex vivo with a Cpfl and a gRNA that targets a dystrophin splice site. In some embodiments, the cells are contacted with one or more nucleic acids encoding the Cpfl and the guide RNA. In some embodiments, the one or more nucleic acids are introduced into the cells using, for example, lipofection or electroporation. Gene editing may also be performed in zygotes. In embodiments, zygotes may be injected with one or more nucleic acids encoding Cpfl and a gRNA that targets a dystrophin splice site. The zygotes may subsequently be injected into a host.

In embodiments, the Cpfl is provided on a vector. In embodiments, the vector contains a Cpfl sequence derived from a Lachnospiraceae bacterium. See, for example, Uniprot Accession No. A0A182DWE3; SEQ ID NO. 76. In embodiments, the vector contains a Cpfl sequence derived from an Acidaminococcus bacterium. See, for example, Uniprot Accession No. U2UMQ6; SEQ ID NO. 75. In some embodiments, the Cpfl sequence is codon optimized for expression in human cells or mouse cells. In some embodiments, the vector further contains a sequence encoding a fluorescent protein, such as GFP, which allows Cpfl -expressing cells to be sorted using fluorescence activated cell sorting (FACS). In some embodiments, the vector is a viral vector such as an adeno-associated viral vector.

In embodiments, the gRNA is provided on a vector. In some embodiments, the vector is a viral vector such as an adeno-associated viral vector. In embodiments, the Cpfl and the guide RNA are provided on the same vector. In embodiments, the Cpfl and the guide RNA are provided on different vectors.

In some embodiments, the cells are additionally contacted with a single-stranded DMD oligonucleotide to effect homology directed repair. In some embodiments, small INDELs restore the protein reading frame of dystrophin ("refraining" strategy). When the refraining strategy is used, the cells may be contacted with a single gRNA. In embodiments, a splice donor or splice acceptor site is disrupted, which results in exon skipping and restoration of the protein reading frame ("exon skipping" strategy). When the exon skipping strategy is used, the cells may be contacted with two or more gRNAs.

Efficiency of in vitro or ex vivo Cpfl -mediated DNA cleavage may be assessed using techniques known to those of skill in the art, such as the T7 El assay. Restoration of DMD expression may be confirmed using techniques known to those of skill in the art, such as RT- PCR, western blotting, and immunocytochemistry.

In some embodiments, in vitro or ex vivo gene editing is performed in a muscle or satellite cell. In some embodiments, gene editing is performed in iPSC or iCM cells. In embodiments, the iPSC cells are differentiated after gene editing. For example, the iPSC cells may be differentiated into a muscle cell or a satellite cell after editing. In embodiments, the iPSC cells are differentiated into cardiac muscle cells, skeletal muscle cells, or smooth muscle cells. In embodiments, the iPSC cells are differentiated into cardiomyocytes. iPSC cells may be induced to differentiate according to methods known to those of skill in the art.

In some embodiments, contacting the cell with the Cpfl and the gRNA restores dystrophin expression. In embodiments, cells which have been edited in vitro or ex vivo, or cells derived therefrom, show levels of dystrophin protein that is comparable to wild type cells. In embodiments, the edited cells, or cells derived therefrom, express dystrophin at a level that is 50%, 60%, 70%, 80%, 90%, 95% or any percentage in between of wild type dystrophin expression levels. In embodiments, the cells which have been edited in vitro or ex vivo, or cells derived therefrom, have a mitochondrial number that is comparable to that of wild type cells. In embodiments the edited cells, or cells derived therefrom, have 50%, 60%, 70%, 80%, 90%, 95% or any percentage in between as many mitochondria as wild type cells. In embodiments, the edited cells, or cells derived therefrom, show an increase in oxygen consumption rate (OCR) compared to non-edited cells at baseline.

C. RNA Pol III and Pol III Promoters

In eukaryotes, RNA polymerase III (also called Pol III) transcribes DNA to synthesize ribosomal 5S rRNA, tRNA and other small RNAs. The genes transcribed by RNA Pol III fall in the category of "housekeeping" genes whose expression is required in all cell types and most environmental conditions. Therefore, the regulation of Pol III transcription is primarily tied to the regulation of cell growth and the cell cycle, thus requiring fewer regulatory proteins than RNA polymerase II. Under stress conditions however, the protein Mafl represses Pol III activity.

In the process of transcription (by any polymerase) there are three main stages: (i) initiation, requiring construction of the RNA polymerase complex on the gene's promoter; (ii) elongation, the synthesis of the RNA transcript; and (iii) termination, the finishing of RNA transcription and disassembly of the RNA polymerase complex.

Promoters under the control of RNA Pol III include those for ribosomal 5S rRNA, tRNA and few other small RNAs such as U6 spliceosomal RNA, RNase P and RNase MRP RNA, 7SL RNA (the RNA component of the signal recognition particles), Vault RNAs, Y RNA, SINEs (short interspersed repetitive elements), 7SK RNA, two microRNAs, several small nucleolar RNAs and several few regulatory antisense RNAs

III. Nucleic Acid Delivery

In some embodiments, one or more nucleic acids (e.g. , an expression vector) are delivered to a cell. The cell may be a mammalian cell, for example a human cell, a mouse cell, or a dog cell. In some embodiments, the cell is an oocyte. In some embodiments, the cell is a non-human oocyte. In some embodiments the cell is a stem cell, such as an iPSC.

As discussed above, in certain embodiments, expression cassettes are employed to express a transcription factor product, either for subsequent purification and delivery to a cell/subject, or for use directly in a genetic-based delivery approach. Provided herein are expression vectors which contain one or more nucleic acids encoding Cpfl and at least one DMD guide RNA that targets a dystrophin splice site. In some embodiments, a nucleic acid encoding Cpf 1 and a nucleic acid encoding at least one guide RNA are provided on the same vector. In further embodiments, a nucleic acid encoding Cpfl and a nucleic acid encoding least one guide RNA are provided on separate vectors.

Expression requires that appropriate signals be provided in the vectors, and include various regulatory elements such as enhancers/promoters from both viral and mammalian sources that drive expression of the genes of interest in cells. Elements designed to optimize messenger RNA stability and translatability in host cells also are defined. The conditions for the use of a number of dominant drug selection markers for establishing permanent, stable cell clones expressing the products are also provided, as is an element that links expression of the drug selection markers to expression of the polypeptide.

A. Regulatory Elements

Throughout this application, the term "expression cassette" is meant to include any type of genetic construct containing a nucleic acid coding for a gene product in which part or all of the nucleic acid encoding sequence is capable of being transcribed and translated, i. e. , is under the control of a promoter. A "promoter" refers to a DNA sequence recognized by the synthetic machinery of the cell, or introduced synthetic machinery, required to initiate the specific transcription of a gene. The phrase "under transcriptional control" means that the promoter is in the correct location and orientation in relation to the nucleic acid to control RNA polymerase initiation and expression of the gene. An "expression vector" is meant to include expression cassettes comprised in a genetic construct that is capable of replication, and thus including one or more of origins of replication, transcription termination signals, poly-A regions, selectable markers, and multipurpose cloning sites.

The term promoter will be used here to refer to a group of transcriptional control modules that are clustered around the initiation site for RNA polymerase II. Much of the thinking about how promoters are organized derives from analyses of several viral promoters, including those for the HSV thymidine kinase (tk) and SV40 early transcription units. These studies, augmented by more recent work, have shown that promoters are composed of discrete functional modules, each consisting of approximately 7-20 bp of DNA, and containing one or more recognition sites for transcriptional activator or repressor proteins.

At least one module in each promoter functions to position the start site for RNA synthesis. The best known example of this is the TATA box, but in some promoters lacking a TATA box, such as the promoter for the mammalian terminal deoxynucleotidyl transferase gene and the promoter for the SV40 late genes, a discrete element overlying the start site itself helps to fix the place of initiation.

In some embodiments, the Cpfl or Cas9 constructs of the disclosure are expressed by a muscle-cell specific promoter. This muscle-cell specific promoter may be constitutively active or may be an inducible promoter.

Additional promoter elements regulate the frequency of transcriptional initiation. Typically, these are located in the region 30-110 bp upstream of the start site, although a number of promoters have recently been shown to contain functional elements downstream of the start site as well. The spacing between promoter elements frequently is flexible, so that promoter function is preserved when elements are inverted or moved relative to one another. In the tk promoter, the spacing between promoter elements can be increased to 50 bp apart before activity begins to decline. Depending on the promoter, it appears that individual elements can function either co-operatively or independently to activate transcription.

In certain embodiments, viral promoters such as the human cytomegalovirus (CMV) immediate early gene promoter, the SV40 early promoter, the Rous sarcoma virus long terminal repeat, rat insulin promoter and glyceraldehyde-3-phosphate dehydrogenase can be used to obtain high-level expression of the coding sequence of interest. The use of other viral or mammalian cellular or bacterial phage promoters which are well-known in the art to achieve expression of a coding sequence of interest is contemplated as well, provided that the levels of expression are sufficient for a given purpose. By employing a promoter with well-known properties, the level and pattern of expression of the protein of interest following transfection or transformation can be optimized. Further, selection of a promoter that is regulated in response to specific physiologic signals can permit inducible expression of the gene product.

Enhancers are genetic elements that increase transcription from a promoter located at a distant position on the same molecule of DNA. Enhancers are organized much like promoters. That is, they are composed of many individual elements, each of which binds to one or more transcriptional proteins. The basic distinction between enhancers and promoters is operational. An enhancer region as a whole must be able to stimulate transcription at a distance; this need not be true of a promoter region or its component elements. On the other hand, a promoter must have one or more elements that direct initiation of RNA synthesis at a particular site and in a particular orientation, whereas enhancers lack these specificities. Promoters and enhancers are often overlapping and contiguous, often seeming to have a very similar modular organization. Below is a list of promoters/enhancers and inducible promoters/enhancers that could be used in combination with the nucleic acid encoding a gene of interest in an expression construct. Additionally, any promoter/enhancer combination (as per the Eukaryotic Promoter Data Base EPDB) could also be used to drive expression of the gene. Eukaryotic cells can support cytoplasmic transcription from certain bacterial promoters if the appropriate bacterial polymerase is provided, either as part of the delivery complex or as an additional genetic expression construct.

The promoter and/or enhancer may be, for example, immunoglobulin light chain, immunoglobulin heavy chain, T-cell receptor, HLA DQ a and/or DQ β, β-interferon, interleukin-2, interleukin-2 receptor, MHC class II 5, MHC class II HLA-Dra, β-Actin, muscle creatine kinase (MCK), prealbumin (transthyretin), elastase I, metallothionein (MTII), collagenase, albumin, a -fetoprotein, t-globin, β-globin, c-fos, c-HA-ras, insulin, neural cell adhesion molecule (NCAM), ai-antitrypain, H2B (TH2B) histone, mouse and/or type I collagen, glucose-regulated proteins (GRP94 and GRP78), rat growth hormone, human serum amyloid A (SAA), troponin I (TN I), platelet-derived growth factor (PDGF), duchenne muscular dystrophy, SV40, polyoma, retroviruses, papilloma virus, hepatitis B virus, human immunodeficiency virus, cytomegalovirus (CMV), and gibbon ape leukemia virus.

The promoter and/or enhancer may be, for example, immunoglobulin light chain, immunoglobulin heavy chain, T-cell receptor, HLA DQ a and/or DQ β, β-interferon, interleukin-2, interleukin-2 receptor, MHC class II 5, MHC class II HLA-Dra, β-Actin, muscle creatine kinase (MCK), prealbumin (transthyretin), elastase I, metallothionein (MTII), collagenase, albumin, a -fetoprotein, t-globin, β-globin, c-fos, c-HA-ras, insulin, neural cell adhesion molecule (NCAM), ai-antitrypain, H2B (TH2B) histone, mouse and/or type I collagen, glucose-regulated proteins (GRP94 and GRP78), rat growth hormone, human serum amyloid A (SAA), troponin I (TN I), platelet-derived growth factor (PDGF), duchenne muscular dystrophy, SV40, polyoma, retroviruses, papilloma virus, hepatitis B virus, human immunodeficiency virus, cytomegalovirus (CMV), and gibbon ape leukemia virus.

In some embodiments, inducible elements may be used. In some embodiments, the inducible element is, for example, MTII, MMTV (mouse mammary tumor virus), β-interferon, adenovirus 5 E2, collagenase, stromelysin, SV40, murine MX gene, GRP78 gene, a-2- macroglobulin, vimentin, MHC class I gene Η-2κ±>, HSP70, proliferin, tumor necrosis factor, and/or thyroid stimulating hormone a gene. In some embodiments, the inducer is phorbol ester (TFA), heavy metals, glucocorticoids, poly(rI)x, poly(rc), E1A, phorbol ester (TPA), interferon, Newcastle Disease Virus, A23187, IL-6, serum, interferon, SV40 large T antigen, PMA, and/or thyroid hormone. Any of the inducible elements described herein may be used with any of the inducers described herein.

Of particular interest are muscle specific promoters. These include the myosin light chain-2 promoter, the a-actin promoter, the troponin 1 promoter; the Na ⁺ /Ca ²⁺ exchanger promoter, the dystrophin promoter, the a7 integrin promoter, the brain natriuretic peptide promoter and the aB-crystallin/small heat shock protein promoter, a-myosin heavy chain promoter and the ANF promoter. In some embodiments, the muscle specific promoter is the CK8 promoter. The CK8 promoter has the following sequence (SEQ ID NO. 77):

CTAGACTAGC ATGCTGCCCA TGTAAGGAGG CAAGGCCTGG GGACACCCGA

GATGCCTGGT

61 TATAATTAAC CCAGACATGT GGCTGCCCCC CCCCCCCCAA CACCTGCTGC

CTCTAAAAAT

12 1 AACCCTGCAT GCCATGTTCC CGGCGAAGGG CCAGCTGTCC CCCGCCAGCT

AGACTCAGCA

18 1 CTTAGTTTAG GAACCAGTGA GCAAGTCAGC CCTTGGGGCA GCCCATACAA

GGCCATGGGG

24 1 CTGGGCAAGC TGCACGCCTG GGTCCGGGGT GGGCACGGTG CCCGGGCAAC

GAGCTGAAAG

301 CTCATCTGCT CTCAGGGGCC CCTCCCTGGG GACAGCCCCT CCTGGCTAGT

CACACCCTGT

361 AGGCTCCTCT ATATAACCCA GGGGCACAGG GGCTGCCCTC ATTCTACCAC

CACCTCCACA

42 1 GCACAGACAG ACACTCAGGA GCCAGCCAGC

In some embodiments, the muscle-cell cell specific promoter is a variant of the CK8 promoter, called CK8e. The CK8e promoter has the following sequence (SEQ ID NO. 78):

1 TGCCCATGTA AGGAGGCAAG GCCTGGGGAC ACCCGAGATG CCTGGTTATA ATTAACCCAG

61 ACATGTGGCT GCCCCCCCCC CCCCAACACC TGCTGCCTCT AAAAATAACC CTGCATGCCA

121 TGTTCCCGGC GAAGGGCCAG CTGTCCCCCG CCAGCTAGAC TCAGCACTTA GTTTAGGAAC

1 81 CAGTGAGCAA GTCAGCCCTT GGGGCAGCCC ATACAAGGCC ATGGGGCTGG GCAAGCTGCA

2 41 CGCCTGGGTC CGGGGTGGGC ACGGTGCCCG GGCAACGAGC TGAAAGCTCA TCTGCTCTCA

3 01 GGGGCCCCTC CCTGGGGACA GCCCCTCCTG GCTAGTCACA CCCTGTAGGC TCCTCTATAT 3 61 AACCCAGGGG CACAGGGGCT GCCCTCATTC TACCACCACC TCCACAGCAC AGACAGACAC 421 TCAGGAGCCA GCCAGC Where a cDNA insert is employed, one will typically desire to include a polyadenylation signal to effect proper polyadenylation of the gene transcript. Any polyadenylation sequence may be employed such as human growth hormone and SV40 polyadenylation signals. Also contemplated as an element of the expression cassette is a terminator. These elements can serve to enhance message levels and to minimize read through from the cassette into other sequences.

B. 2A Protease

The inventor utilizes the 2A-like self-cleaving domain from the insect virus Thosea asigna (TaV 2A peptide; SEQ ID NO. 79; EGRGSLLTCGDVEENPGP). These 2A-like domains have been shown to function across Eukaryotes and cause cleavage of amino acids to occur co-translationally within the 2A-like peptide domain. Therefore, inclusion of TaV 2A peptide allows the expression of multiple proteins from a single mRNA transcript. Importantly, the domain of TaV when tested in eukaryotic systems has shown greater than 99% cleavage activity. Other acceptable 2A-like peptides include, but are not limited to, equine rhinitis A virus (ERAV) 2A peptide (SEQ ID NO. 80; QCTNYALLKLAGDVESNPGP), porcine tescho virus- 1 (PTVl) 2A peptide (SEQ ID NO. 81; ATNFSLLKQAGDVEENPGP) and foot and mouth disease virus (FMDV) 2A peptide (SEQ ID NO. 82; PVKQLLNFDLLKLAGDVESNPGP) or modified versions thereof.

In some embodiments, the 2A peptide is used to express a reporter and a Cfpl or a Cas9 simultaneously. The reporter may be, for example, GFP.

Other self-cleaving peptides that may be used include, but are not limited to nuclear inclusion protein a (Nia) protease, a P 1 protease, a 3C protease, a L protease, a 3C-like protease, or modified versions thereof.

C. Delivery of Expression Vectors

There are a number of ways in which expression vectors may be introduced into cells. In certain embodiments, the expression construct comprises a virus or engineered construct derived from a viral genome. The ability of certain viruses to enter cells via receptor-mediated endocytosis, to integrate into host cell genome and express viral genes stably and efficiently have made them attractive candidates for the transfer of foreign genes into mammalian cells. These have a relatively low capacity for foreign DNA sequences and have a restricted host spectrum. Furthermore, their oncogenic potential and cytopathic effects in permissive cells raise safety concerns. They can accommodate only up to 8 kB of foreign genetic material but can be readily introduced in a variety of cell lines and laboratory animals.

One of the preferred methods for in vivo delivery involves the use of an adenovirus expression vector. "Adenovirus expression vector" is meant to include those constructs containing adenovirus sequences sufficient to (a) support packaging of the construct and (b) to express an antisense polynucleotide that has been cloned therein. In this context, expression does not require that the gene product be synthesized.

The expression vector comprises a genetically engineered form of adenovirus. Knowledge of the genetic organization of adenovirus, a 36 kB, linear, double-stranded DNA virus, allows substitution of large pieces of adenoviral DNA with foreign sequences up to 7 kB. In contrast to retrovirus, the adenoviral infection of host cells does not result in chromosomal integration because adenoviral DNA can replicate in an episomal manner without potential genotoxicity. Also, adenoviruses are structurally stable, and no genome rearrangement has been detected after extensive amplification. Adenovirus can infect virtually all epithelial cells regardless of their cell cycle stage. So far, adenoviral infection appears to be linked only to mild disease such as acute respiratory disease in humans.

Adenovirus is particularly suitable for use as a gene transfer vector because of its midsized genome, ease of manipulation, high titer, wide target cell range and high infectivity. Both ends of the viral genome contain 100-200 base pair inverted repeats (ITRs), which are cis elements necessary for viral DNA replication and packaging. The early (E) and late (L) regions of the genome contain different transcription units that are divided by the onset of viral DNA replication. The El region (El A and EIB) encodes proteins responsible for the regulation of transcription of the viral genome and a few cellular genes. The expression of the E2 region (E2A and E2B) results in the synthesis of the proteins for viral DNA replication. These proteins are involved in DNA replication, late gene expression and host cell shut-off The products of the late genes, including the majority of the viral capsid proteins, are expressed only after significant processing of a single primary transcript issued by the major late promoter (MLP). The MLP, (located at 16.8 m.u.) is particularly efficient during the late phase of infection, and all the mRNAs issued from this promoter possess a 5'-tripartite leader (TPL) sequence which makes them preferred mRNAs for translation.

In one system, recombinant adenovirus is generated from homologous recombination between shuttle vector and provirus vector. Due to the possible recombination between two proviral vectors, wild-type adenovirus may be generated from this process. Therefore, it is critical to isolate a single clone of virus from an individual plaque and examine its genomic structure.

Generation and propagation of the current adenovirus vectors, which are replication deficient, depend on a unique helper cell line, designated 293, which was transformed from human embryonic kidney cells by Ad5 DNA fragments and constitutively expresses El proteins. Since the E3 region is dispensable from the adenovirus genome, the current adenovirus vectors, with the help of 293 cells, carry foreign DNA in either the El, the D3 or both regions. In nature, adenovirus can package approximately 105% of the wild-type genome, providing capacity for about 2 extra kb of DNA. Combined with the approximately 5.5 kb of DNA that is replaceable in the El and E3 regions, the maximum capacity of the current adenovirus vector is under 7.5 kb, or about 15% of the total length of the vector. More than 80% of the adenovirus viral genome remains in the vector backbone and is the source of vector- borne cytotoxicity. Also, the replication deficiency of the El-deleted virus is incomplete.

Helper cell lines may be derived from human cells such as human embryonic kidney cells, muscle cells, hematopoietic cells or other human embryonic mesenchymal or epithelial cells. Alternatively, the helper cells may be derived from the cells of other mammalian species that are permissive for human adenovirus. Such cells include, e.g. , Vero cells or other monkey embryonic mesenchymal or epithelial cells. As stated above, the preferred helper cell line is 293.

Methods for culturing 293 cells and propagating adenovirus are known to those of skill in the art. In one format, natural cell aggregates are grown by inoculating individual cells into 1 liter siliconized spinner flasks (Techne, Cambridge, UK) containing 100-200 ml of medium. Following stirring at 40 rpm, the cell viability is estimated with trypan blue. In another format, Fibra-Cel microcarriers (Bibby Sterlin, Stone, UK) (5 g/1) is employed as follows. A cell inoculum, resuspended in 5 ml of medium, is added to the carrier (50 ml) in a 250 ml Erlenmeyer flask and left stationary, with occasional agitation, for 1 to 4 h. The medium is then replaced with 50 ml of fresh medium and shaking initiated. For virus production, cells are allowed to grow to about 80% confluence, after which time the medium is replaced (to 25% of the final volume) and adenovirus added at an MOI of 0.05. Cultures are left stationary overnight, following which the volume is increased to 100% and shaking commenced for another 72 h.

The adenoviruses of the disclosure are replication defective, or at least conditionally replication defective. The adenovirus may be of any of the 42 different known serotypes or subgroups A-F. Adenovirus type 5 of subgroup C is the preferred starting material in order to obtain the conditional replication-defective adenovirus vector for use as described herein.

As stated above, the typical vector of the disclosure is replication defective and will not have an adenovirus El region. Thus, it will be most convenient to introduce the polynucleotide encoding the gene of interest at the position from which the El-coding sequences have been removed. However, the position of insertion of the construct within the adenovirus sequences is not critical. The polynucleotide encoding the gene of interest may also be inserted in lieu of the deleted E3 region in E3 replacement vectors, or in the E4 region where a helper cell line or helper virus complements the E4 defect.

Adenovirus is easy to grow and manipulate and exhibits broad host range in vitro and in vivo. This group of viruses can be obtained in high titers, e.g. , 10 ⁹ -10 ¹² plaque-forming units per ml, and they are highly infective. The life cycle of adenovirus does not require integration into the host cell genome. The foreign genes delivered by adenovirus vectors are episomal and, therefore, have low genotoxicity to host cells. No side effects have been reported in studies of vaccination with wild-type adenovirus, demonstrating their safety and therapeutic potential as in vivo gene transfer vectors.

Adenovirus vectors have been used in eukaryotic gene expression and vaccine development. Animal studies suggested that recombinant adenovirus could be used for gene therapy. Studies in administering recombinant adenovirus to different tissues include trachea instillation, muscle injection, peripheral intravenous injections and stereotactic inoculation into the brain.

The retroviruses are a group of single-stranded RNA viruses characterized by an ability to convert their RNA to double-stranded DNA in infected cells by a process of reverse- transcription. The resulting DNA then stably integrates into cellular chromosomes as a provirus and directs synthesis of viral proteins. The integration results in the retention of the viral gene sequences in the recipient cell and its descendants. The retroviral genome contains three genes, gag, pol, and env that code for capsid proteins, polymerase enzyme, and envelope components, respectively. A sequence found upstream from the gag gene contains a signal for packaging of the genome into virions. Two long terminal repeat (LTR) sequences are present at the 5' and 3' ends of the viral genome. These contain strong promoter and enhancer sequences and are also required for integration in the host cell genome.

In order to construct a retroviral vector, a nucleic acid encoding a gene of interest is inserted into the viral genome in the place of certain viral sequences to produce a virus that is replication-defective. In order to produce virions, a packaging cell line containing the gag, pol, and env genes but without the LTR and packaging components is constructed. When a recombinant plasmid containing a cDNA, together with the retroviral LTR and packaging sequences is introduced into this cell line (by calcium phosphate precipitation for example), the packaging sequence allows the RNA transcript of the recombinant plasmid to be packaged into viral particles, which are then secreted into the culture media. The media containing the recombinant retroviruses is then collected, optionally concentrated, and used for gene transfer. Retroviral vectors are able to infect a broad variety of cell types. However, integration and stable expression require the division of host cells.

A novel approach designed to allow specific targeting of retrovirus vectors was recently developed based on the chemical modification of a retrovirus by the chemical addition of lactose residues to the viral envelope. This modification could permit the specific infection of hepatocytes via sialoglycoprotein receptors.

A different approach to targeting of recombinant retroviruses may be used, in which biotinylated antibodies against a retroviral envelope protein and against a specific cell receptor are used. The antibodies are coupled via the biotin components by using streptavidin. Using antibodies against major histocompatibility complex class I and class II antigens, it has been demonstrated the infection of a variety of human cells that bore those surface antigens with an ecotropic virus in vitro.

There are certain limitations to the use of retrovirus vectors. For example, retrovirus vectors usually integrate into random sites in the cell genome. This can lead to insertional mutagenesis through the interruption of host genes or through the insertion of viral regulatory sequences that can interfere with the function of flanking genes. Another concern with the use of defective retrovirus vectors is the potential appearance of wild-type replication-competent virus in the packaging cells. This can result from recombination events in which the intact- sequence from the recombinant virus inserts upstream from the gag, pol, env sequence integrated in the host cell genome. However, new packaging cell lines are now available that should greatly decrease the likelihood of recombination.

Other viral vectors may be employed as expression constructs. Vectors derived from viruses such as vaccinia virus, adeno-associated virus (AAV), and herpesviruses may be employed. They offer several attractive features for various mammalian cells.

In embodiments, the AAV vector is replication-defective or conditionally replication defective. In embodiments, the AAV vector is a recombinant AAV vector. In some embodiments, the AAV vector comprises a sequence isolated or derived from an AAV vector of serotype AAV1, AAV2, AAV3, AAV4, AAV5, AAV6, AAV7, AAV 8, AAV 9, AAV 10, AAV11, or any combination thereof. In some embodiments, the AAV vector is not an AAV9 vector. In some embodiments, the AAV vector is an AAVRh.74 vector.

In some embodiments, a single viral vector is used to deliver a nucleic acid encoding Cpf 1 or Cas9 and at least one gRNA to a cell. In some embodiments, Cpf 1 or Cas9 is provided to a cell using a first viral vector and at least one gRNA is provided to the cell using a second viral vector. In order to effect expression of sense or antisense gene constructs, the expression construct must be delivered into a cell. The cell may be a muscle cell, a satellite cell, a mesangioblast, a bone marrow derived cell, a stromal cell or a mesenchymal stem cell. In embodiments, the cell is a cardiac muscle cell, a skeletal muscle cell, or a smooth muscle cell. In embodiments, the cell is a cell in the tibialis anterior, quadriceps, soleus, diaphragm or heart. In some embodiments, the cell is an induced pluripotent stem cell (iPSC) or inner cell mass cell (iCM). In further embodiments, the cell is a human iPSC or a human iCM. In some embodiments, human iPSCs or human iCMs of the disclosure may be derived from a cultured stem cell line, an adult stem cell, a placental stem cell, or from another source of adult or embryonic stem cells that does not require the destruction of a human embryo. Delivery to a cell may be accomplished in vitro, as in laboratory procedures for transforming cells lines, or in vivo or ex vivo, as in the treatment of certain disease states. One mechanism for delivery is via viral infection where the expression construct is encapsidated in an infectious viral particle.

Several non-viral methods for the transfer of expression constructs into cultured mammalian cells also are contemplated by the present disclosure. These include calcium phosphate precipitation, DEAE-dextran, electroporation, direct microinjection, DNA-loaded liposomes, and lipofectamine-DNA complexes, cell sonication, gene bombardment using high velocity microprojectiles, and receptor-mediated transfection. Some of these techniques may be successfully adapted for in vivo or ex vivo use.

Once the expression construct has been delivered into the cell the nucleic acid encoding the gene of interest may be positioned and expressed at different sites. In certain embodiments, the nucleic acid encoding the gene may be stably integrated into the genome of the cell. This integration may be in the cognate location and orientation via homologous recombination (gene replacement) or it may be integrated in a random, non-specific location (gene augmentation). In yet further embodiments, the nucleic acid may be stably maintained in the cell as a separate, episomal segment of DNA. Such nucleic acid segments or "episomes" encode sequences sufficient to permit maintenance and replication independent of or in synchronization with the host cell cycle. How the expression construct is delivered to a cell and where in the cell the nucleic acid remains is dependent on the type of expression construct employed.

In yet another embodiment, the expression construct may simply consist of naked recombinant DNA or plasmids. Transfer of the construct may be performed by any of the methods mentioned above which physically or chemically permeabilize the cell membrane. This is particularly applicable for transfer in vitro but it may be applied to in vivo use as well. One group successfully injected polyomavirus DNA in the form of calcium phosphate precipitates into liver and spleen of adult and newborn mice demonstrating active viral replication and acute infection. Another group also demonstrated that direct intraperitoneal injection of calcium phosphate-precipitated plasmids results in expression of the transfected genes. DNA encoding a gene of interest may also be transferred in a similar manner in vivo and express the gene product.

In still another embodiment for transferring a naked DNA expression construct into cells may involve particle bombardment. This method depends on the ability to accelerate DNA-coated microprojectiles to a high velocity allowing them to pierce cell membranes and enter cells without killing them. Several devices for accelerating small particles have been developed. One such device relies on a high voltage discharge to generate an electrical current, which in turn provides the motive force. The microprojectiles used have consisted of biologically inert substances such as tungsten or gold beads.

In some embodiments, the expression construct is delivered directly to the liver, skin, and/or muscle tissue of a subject. This may require surgical exposure of the tissue or cells, to eliminate any intervening tissue between the gun and the target organ, i.e. , ex vivo treatment. Again, DNA encoding a particular gene may be delivered via this method and still be incorporated by the present disclosure.

In a further embodiment, the expression construct may be entrapped in a liposome.

Liposomes are vesicular structures characterized by a phospholipid bilayer membrane and an inner aqueous medium. Multilamellar liposomes have multiple lipid layers separated by aqueous medium. They form spontaneously when phospholipids are suspended in an excess of aqueous solution. The lipid components undergo self-rearrangement before the formation of closed structures and entrap water and dissolved solutes between the lipid bilayers. Also contemplated are lipofectamine-DNA complexes.

Liposome-mediated nucleic acid delivery and expression of foreign DNA in vitro has been very successful. A reagent known as Lipofectamine 2000™ is widely used and commercially available. In certain embodiments, the liposome may be complexed with ahemagglutinating virus (HVJ), to facilitate fusion with the cell membrane and promote cell entry of liposome- encapsulated DNA. In other embodiments, the liposome may be complexed or employed in conjunction with nuclear non-histone chromosomal proteins (HMG-1). In yet further embodiments, the liposome may be complexed or employed in conjunction with both HVJ and HMG-1. In that such expression constructs have been successfully employed in transfer and expression of nucleic acid in vitro and in vivo, then they are applicable for the present disclosure. Where a bacterial promoter is employed in the DNA construct, it also will be desirable to include within the liposome an appropriate bacterial polymerase.

Other expression constructs which can be employed to deliver a nucleic acid encoding a particular gene into cells are receptor-mediated delivery vehicles. These take advantage of the selective uptake of macromolecules by receptor-mediated endocytosis in almost all eukaryotic cells. Because of the cell type-specific distribution of various receptors, the delivery can be highly specific.

Receptor-mediated gene targeting vehicles generally consist of two components: a cell receptor-specific ligand and a DNA-binding agent. Several ligands have been used for receptor-mediated gene transfer. The most extensively characterized ligands are asialoorosomucoid (ASOR) and transferrin. A synthetic neoglycoprotein, which recognizes the same receptor as ASOR, has been used as a gene delivery vehicle and epidermal growth factor (EGF) has also been used to deliver genes to squamous carcinoma cells.

IV. Pharmaceutical Compositions and Delivery Methods

For clinical applications, pharmaceutical compositions are prepared in a form appropriate for the intended application. Generally, this entails preparing compositions that are essentially free of pyrogens, as well as other impurities that could be harmful to humans or animals.

Appropriate salts and buffers are used to render drugs, proteins or delivery vectors stable and allow for uptake by target cells. Aqueous compositions of the present disclosure comprise an effective amount of the drug, vector or proteins, dissolved or dispersed in a pharmaceutically acceptable carrier or aqueous medium. The phrase "pharmaceutically or pharmacologically acceptable" refer to molecular entities and compositions that do not produce adverse, allergic, or other untoward reactions when administered to an animal or a human. As used herein, "pharmaceutically acceptable carrier" includes solvents, buffers, solutions, dispersion media, coatings, antibacterial and antifungal agents, isotonic and absorption delaying agents and the like acceptable for use in formulating pharmaceuticals, such as pharmaceuticals suitable for administration to humans. The use of such media and agents for pharmaceutically active substances is well known in the art. Any conventional media or agent that is not incompatible with the active ingredients described herein, its use in therapeutic compositions may be used. Supplementary active ingredients also can be incorporated into the compositions, provided they do not inactivate the vectors or cells of the compositions.

In some embodiments, the active compositions of the present disclosure may include classic pharmaceutical preparations. Administration of these compositions according to the present disclosure may be via any common route so long as the target tissue is available via that route, but generally including systemic administration. This includes oral, nasal, or buccal. Alternatively, administration may be by intradermal, subcutaneous, intramuscular, intraperitoneal or intravenous injection, or by direct injection into muscle tissue. Such compositions are normally administered as pharmaceutically acceptable compositions, as described supra.

The active compounds may also be administered parenterally or intraperitoneally. By way of illustration, solutions of the active compounds as free base or pharmacologically acceptable salts can be prepared in water suitably mixed with a surfactant, such as hydroxypropylcellulose. Dispersions can also be prepared in glycerol, liquid polyethylene glycols, and mixtures thereof and in oils. Under ordinary conditions of storage and use, these preparations generally contain a preservative to prevent the growth of microorganisms.

The pharmaceutical forms suitable for injectable use include, for example, sterile aqueous solutions or dispersions and sterile powders for the extemporaneous preparation of sterile injectable solutions or dispersions. Generally, these preparations are sterile and fluid to the extent that easy injectability exists. Preparations should be stable under the conditions of manufacture and storage and should be preserved against the contaminating action of microorganisms, such as bacteria and fungi. Appropriate solvents or dispersion media may contain, for example, water, ethanol, polyol (for example, glycerol, propylene glycol, and liquid polyethylene glycol, and the like), suitable mixtures thereof, and vegetable oils. The proper fluidity can be maintained, for example, by the use of a coating, such as lecithin, by the maintenance of the required particle size in the case of dispersion and by the use of surfactants. The prevention of the action of microorganisms can be brought about by various antibacterial and antifungal agents, for example, parabens, chlorobutanol, phenol, sorbic acid, thimerosal, and the like. In many cases, it will be preferable to include isotonic agents, for example, sugars or sodium chloride. Prolonged absorption of the injectable compositions can be brought about by the use in the compositions of agents delaying absorption, for example, aluminum monostearate and gelatin.

Sterile injectable solutions may be prepared by incorporating the active compounds in an appropriate amount into a solvent along with any other ingredients (for example as enumerated above) as desired, followed by filtered sterilization. Generally, dispersions are prepared by incorporating the various sterilized active ingredients into a sterile vehicle which contains the basic dispersion medium and the desired other ingredients, e.g. , as enumerated above. In the case of sterile powders for the preparation of sterile injectable solutions, the preferred methods of preparation include vacuum-drying and freeze-drying techniques which yield a powder of the active ingredient(s) plus any additional desired ingredient from a previously sterile-filtered solution thereof.

In some embodiments, the compositions of the present disclosure are formulated in a neutral or salt form. Pharmaceutically-acceptable salts include, for example, acid addition salts (formed with the free amino groups of the protein) derived from inorganic acids (e.g. , hydrochloric or phosphoric acids, or from organic acids (e.g. , acetic, oxalic, tartaric, mandelic, and the like). Salts formed with the free carboxyl groups of the protein can also be derived from inorganic bases (e.g. , sodium, potassium, ammonium, calcium, or ferric hydroxides) or from organic bases (e.g. , isopropylamine, trimethylamine, histidine, procaine and the like.)

Upon formulation, solutions are preferably administered in a manner compatible with the dosage formulation and in such amount as is therapeutically effective. The formulations may easily be administered in a variety of dosage forms such as injectable solutions, drug release capsules and the like. For parenteral administration in an aqueous solution, for example, the solution generally is suitably buffered and the liquid diluent first rendered isotonic for example with sufficient saline or glucose. Such aqueous solutions may be used, for example, for intravenous, intramuscular, subcutaneous and intraperitoneal administration. Preferably, sterile aqueous media are employed as is known to those of skill in the art, particularly in light of the present disclosure. By way of illustration, a single dose may be dissolved in 1 ml of isotonic NaCl solution and either added to 1000 ml of hypodermoclysis fluid or injected at the proposed site of infusion. Some variation in dosage will necessarily occur depending on the condition of the subject being treated. The person responsible for administration will, in any event, determine the appropriate dose for the individual subject. Moreover, for human administration, preparations should meet sterility, pyrogenicity, general safety and purity standards as required by FDA Office of Biologies standards. In some embodiments, the Cpf 1 or Cas9 and gRNAs described herein may be delivered to the patient using adoptive cell transfer (ACT). In adoptive cell transfer, one or more expression constructs are provided ex vivo to cells which have originated from the patient (autologous) or from one or more individual(s) other than the patient (allogeneic). The cells are subsequently introduced or reintroduced into the patient. Thus, in some embodiments, one or more nucleic acids encoding Cpfl or Cas9 and a guide RNA that targets a dystrophin splice site are provided to a cell ex vivo before the cell is introduced or reintroduced to a patient.

V. DMD Myoediting

A dystrophin mutation that disrupts the actin-binding domain may be generated. The inventors generated a human iPSC line with a deletion of exons 8 and 9 in the DMD gene using CRISPR/Cas9-mediated editing to analyze the effect ABD-1 deletions on muscle function (FIG. 1A). Two single guide RNAs (gRNAs), targeting sequences in intron 7 and intron 9 of the DMD gene in the presence of Streptococcus pyogenes Cas9, were used to delete DMD exons 8-9 in healthy human iPSCs (FIG. IB). This induced DMD mutant human cell line is referred to as ΔΕχ8-9 iDMD, and represents the common 5 '-proximal hot spot mutations. Human iPSCs harboring DMD exons 8-9 deletions were picked by clonal selection and the mutation was confirmed by sequencing genomic DNA (FIG. 1C). The human ΔΕχ8-9 iDMD iPSC line has a 1,909 bp deletion, which generates a newly formed junction between intron 7 to intron 9 (FIG. 1C). Primers within intron 7 flanking the gRNA-7 targeting site were used to validate the deletion and as expected showed no PCR product with ΔΕχ8-9 iDMD compared to control iPSC with a 424 bp product (FIG. 1C). The AEx8-9 iDMD iPSCs were differentiated to cardiomyocytes and RT-PCR was performed using forward and reverse primers targeting exons 1 and 10, respectively (FIG. ID). Sequencing of the RT-PCR products showed splicing of exon 7 to exon 10, which created a premature stop codon following the first 5 amino acids encoded by exon 10 (FIG. ID). Loss of dystrophin protein was confirmed by Western blot analysis and immunocytochemistry in ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes (FIGS. 1E-F).

The ΔΕχ8-9 iDMD mutation may be corrected by exon deletion strategies. For example, three different CRISPR/Cas9-mediated strategies were used to correct the mutation in ΔΕχ8-9 iDMD iPSCs in order to restore the dystrophin open reading frame (FIG. 2A). These strategies generated dystrophin with different truncations in the ABD-1 domain, providing a means of analyzing the effect of ABD-1 deletions on muscle function. For each strategy, the inventors applied two gRNAs to delete multiple exons and analyzed two independent iPSC clones. Strategy #1 generated ΔΕχ3-9 iPSCs, by targeting intron 2 and intron 7 with gRNAs, and deleting exons 3 through 7 (FIG. 2B). Genomic sequencing of the genome edited region showed a hybrid junction of intron 2 to intron 7, confirming generation of ΔΕχ3-9 iPSCs (FIG. 2C). Following differentiation of ΔΕχ3-9 iPSCs to cardiomyocytes, RT-PCR was performed using primers within exon 1 and exon 10 (FIG. 2D). Sequencing of the RT-PCR product showed splicing of exon 2 to exon 10. Restoration of the DMD open reading frame generated a truncated dystrophin protein lacking the ABS-2 and ABS-3 regions. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry of ΔΕχ3-9 iPSC-derived cardiomyocytes (FIGS. 2E-F; FIG. 7A).

Strategy #2 generated ΔΕχ6-9 iPSCs, using gRNAs targeting intron 5 and intron 7 in order to delete exons 6 and 7 (FIG. 3 A). This form of CRISPR/Cas9 editing resulted in deletion of exon 6 through exon 9, which was confirmed by sequencing of genomic PCR products (FIG. 3B). Following differentiation, RT-PCR using primers within exon 1 and exon 10 was performed on ΔΕχ6-9 iPSC-derived cardiomyocytes and sequencing of the RT-PCR product confirmed splicing of exon 5 to exon 10 (FIG. 3C). Restoration of dystrophin expression generated a truncated protein lacking the ABS-3 region. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry in ΔΕχ6-9 iPSC-derived cardiomyocytes (FIGS. 3D-E; FIGS. 7A-B).

Strategy #3 generated ΔΕχ7-11 iPSCs, using gRNAs targeting introns 6 and 11 to delete exons 7 to 11 of ΔΕχ8-9 iDMD iPSCs (FIG. 4A). This genome editing strategy introduced the largest deletion of 164 kb in ΔΕχ8-9 iDMD iPSCs. Similar to the first two correction strategies, single cell-derived colonies were picked from the corrected iPSC pool and the genomic region was sequenced to confirm editing (FIG. 4B). The ΔΕχ7-11 iPSCs were differentiated to cardiomyocytes, and RT PCR was performed with primers within exon 5 and exon 12 to reveal splicing of exon 6 to exon 12, as seen by sequencing (FIG. 4C). Restoration of the DMD open reading frame of ΔΕχ7-11 iPSC-derived cardiomyocytes generated a truncated dystrophin protein lacking amino acids 177-444 but with all three ABS intact. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry (FIGS. 4D-E; FIGS. 7A-B). The inventors observed less dystrophin expression in ΔΕχ7-11 iPSC-derived cardiomyocytes compared to control iPSC-derived cardiomyocytes and the other corrected lines, ΔΕχ3-9 and ΔΕχ6-9. Previously, it was shown that missense mutations associated with the DMD phenotype caused steady-state decreases in dystrophin protein levels inversely proportional to the tertiary stability and directly caused by proteasomal degradation. Consistent with this report, the inventors observed increased dystrophin protein levels when ΔΕχ7-11 iPSC-derived cardiomyocytes were treated with the proteasome inhibitor MG-132, as measured by Western blot analysis (FIG. 4F; FIG. 7C). This suggests that deletion of DMD exons 7-11, which produces dystrophin lacking 177-444 amino acids, results in a truncated form of dystrophin that is degraded.

Functional restoration of iPSC-derived cardiomyocytes may be achieved by correction of ABD-1 mutations. To assess the consequences of ABD-1 mutations and the effect of correction by genome editing on dystrophin function in muscle, the inventors analyzed spontaneous Ca ²⁺ activity in iPSC-derived cardiomyocytes (FIG. 5A). For each CRISPR/Cas9- edited correction, the inventors analyzed data sets from two corrected clones. As expected, the inventors observed that calcium release and reuptake parameters, including time to peak, Ca ²⁺ decay rate, and transient duration were significantly higher in the ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes compared to isogenic control cells (FIGS. 5B-D). Additionally, greater than 50% of the ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes demonstrated abnormal Ca ²⁺ activity, which represents an arrhythmogenic-like phenotype (FIG. 5E). In contrast, only 10.7% of the control iPSC-derived cardiomyocytes were desynchronized (FIG. 5E).

Corrected ΔΕχ3-9 iPSC-derived cardiomyocytes displayed normalized Ca ²⁺ transient kinetics and decreased asynchronous activity similar to isogenic control cells, suggesting restoration of dystrophin function (FIGS. 5A-G). Cardiomyocytes derived from ΔΕχ6-9 iPSCs showed improved Ca ²⁺ handling, although not to the level seen with ΔΕχ3-9 cells (FIGS. 5A- G). The Ca ²⁺ decay rate and Ca ²⁺ transient duration were shorter in ΔΕχ6-9 iPSC-derived cardiomyocytes (FIGS. 5B and 5D) and the number of arrhythmic cells was decreased to 29% compared to ΔΕχ8-9 iDMD cells (FIG. 5E). However, the parameters observed in the ΔΕχ6-9 iPSC-derived cardiomyocytes did not reach the levels of control iPSC-derived cardiomyocytes. The ΔΕχ7-11 iPSC-derived cardiomyocytes, similarly to ΔΕχ6-9 cells, showed improvement in Ca ²⁺ release and uptake when compared to ΔΕχ8-9 iDMD cardiomyocytes (FIGS. 5B and 5C), however they did not reach control cell levels. iPSC-derived cardiomyocytes with ΔΕχ7- 11 had the highest number of arrhythmic cells, up to 35.9%, among the corrected cell lines (FIG. 5E). Taken together, these findings indicate that deletion of exons 3-9 (ΔΕχ3-9) to correct DMD mutations within the 5 '-proximal hot spot of the DMD gene produces a truncated dystrophin protein with restoration of function.

Further assessment of the functional properties of the iPSC-derived cardiomyocyte cell lines was performed by generating engineered heart muscle (EHM), which displays phenotypic properties of the postnatal myocardium. EHM was generated as shown in FIG. 8A using -95% pure iPSC-derived cardiomyocytes, as measured by a-actinin expression (FIG. 8B). Consistent with the calcium handling data, the inventors observed enhanced contractile performance in all corrected cell lines, with the most pronounced effects in ΔΕχ3-9 EHM (FIG. 5F). Moreover, the inventors observed an increased tendency for arrhythmic contractions in ΔΕχ8-9 iDMD- EHM and a reduction in ΔΕχ3-9 EHM (FIG. 8C).

Correction of DMD patient-derived iPSCs may be achieved by deleting exons 3-9. An iPSC line, referred to as ρΔΕχ3-7, was generated from a DMD patient (SC604A-MD) with a deletion of exons 3-7 in the DMD gene. ρΔΕχ3-7 iPSCs are used as patient-in-a-dish model of DMD with a mutation in the ABD-1. To evaluate the effectiveness of exon 3-9 deletion in restoration of cardiomyocyte function, the inventors corrected ρΔΕχ3-7 by deleting exons 8 and 9 to generate ρΔΕχ3-9. Two gRNAs targeting introns 7 and 9 were used to excise exons 8-9, generating ρΔΕχ3-9 iPSCs (FIG. 6A). Sequencing of genomic DNA PCR products confirmed the hybrid junction of intron 7 to intron 9 in a single cell-derived iPSC colony (FIG. 6B). Both ρΔΕχ3-7 and ρΔΕχ3-9 iPSC lines were differentiated into cardiomyocytes and RT- PCR was performed with primers targeting the 5'-UTR and exon 10. In ρΔΕχ3-7 iPSC-derived cardiomyocytes, the junction of exon 2 to exon 8 was seen by sequencing of the RT-PCR product (FIG. 6C). In corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes, the junction of exon 2 to exon 10 was demonstrated by sequencing the RT-PCR product (FIG. 6C). Western blot analysis showed a low level of dystrophin expression in DMD ρΔΕχ3-7 iPSC-derived cardiomyocytes (FIG. 6D). This is consistent with previous reports showing low levels of dystrophin protein expression in patients lacking exons 3-7 due to re-initiation of translation at an in-frame start codon in exon 8 (Fletcher et al, 2012; Winnard et al, 1995). Corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes showed dystrophin expression similar to normal control cardiomyocytes (FIG. 6D). These findings were confirmed by immunocytochemistry (FIG. 6E).

Spontaneous Ca ²⁺ activity was assessed in these patient-derived cells as a measure of iPSC-derived cardiomyocyte functionality (FIGS. 6F-I). In ρΔΕχ3-7 DMD iPSC-derived cardiomyocytes, Ca ²⁺ time to peak and time to half decay were elevated (FIGS. 6F-G). This caused an overall slower Ca ²⁺ transient (FIG. 6H) and elevated the number of arrhythmic cells, up to 53% (FIG. 61). Corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes had significantly improved time to peak and faster Ca ²⁺ decay (FIGS. 6F-G). Although these parameters of the corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes did not reach normal control levels, the number of arrhythmic cells was 18.5%, similar to control iPSC-derived cardiomyocytes (FIG. 61). These findings show improved iPSC-derived cardiomyocyte function after genomic editing to excise exons 3-9 of the DMD gene in DMD patient-derived iPSCs.

ΔΕχ8-9 DMD mouse models may be used to recapitulate the muscle dystrophy phenotype. To investigate CRISPR/Cas9-mediated exon skipping and refraining in vivo, a mimic of the human ABD-1 region mutations was generated in a mouse model by deleting exons 8 and 9, using CRISPR/Cas9 system directed by two single guide RNAs (sgRNA) (FIG. 9A and Table 2). The inventors designed and validated sgRNAs targeting introns, flanking 5' end and 3' ends of Dmd exons 8 and 9 respectively. C57BL/6 zygotes were co-injected with in vitro transcribed Cas9 mRNA and in vitro transcribed sgRNAs, and then re-implanted into pseudo-pregnant females.

The deletion of Dmd exons 8-9 was confirmed by DNA genotyping. Mice lacking exons 8-9 showed pronounced dystrophic muscle changes in 1 and 2 month-old mice (FIG. 9B). The deletion of these exons placed the dystrophin gene out of frame leading to the absence of dystrophin protein in skeletal muscle and heart (FIG. 9C). The grip strength of the ΔΕχ8-9 DMD was significantly weaker than that in wild-type mice (FIG. 9D). Serum analysis of the ΔΕχ8-9 DMD mice shows a significant increase of creatine kinase (CK) level, which is a sign of muscle damage (FIG. 9E). Taken together, dystrophin protein expression, muscle histology, muscle strength and serum creatine kinase level validated the dystrophic phenotype of the ΔΕχ8-9 DMD mouse model.

Dystrophin protein serves as a muscular shock absorber by providing a structural link between the cytoskeleton and the extracellular matrix to maintain muscle integrity. Essential regions of the dystrophin protein are located at both ends of the protein; the amino-terminus contains ABD-1, the actin binding domain, and the carboxy -terminus includes binding sites for components of the dystrophin associated protein complex, such as sarcoglycans and dystroglycans. Genomic mutations in the DMD gene encoding non-essential regions have been excised by exon skipping and deletion to restore functional, albeit truncated, dystrophin protein. Here, the inventors use CRISPR/Cas9-mediated editing to selectively delete exons 3- 9 of the DMD gene encoding part of the essential ABD-1 region. With precision genomic editing, they can modify the essential amino-terminal region and restore dystrophin expression and cardiomyocyte function.

To provide an isogenic control iPSC line for these studies, the inventors generated an iDMD model (ΔΕχ8-9) by deleting DMD exons 8 and 9 in a healthy donor-derived iPSC line. They confirmed ΔΕχ8-9 iDMD iPSCs as a model of DMD by showing that cardiomyocytes derived from these cells do not express dystrophin protein and display arrhythmias attributable to impaired Ca ²⁺ cycling. The lack of dystrophin results in the loss of linkage between the extracellular matrix and the actin cytoskeleton, destabilizing membrane proteins and resulting in generation of reactive oxygen species and excessive Ca ²⁺ entry. Consistent with these findings, it was shown that cardiomyocytes differentiated from DMD iPSCs have slower Ca ²⁺ cycling. Additionally, others have reported cellular abnormalities such as elevated levels of resting Ca ²⁺ , mitochondrial damage and apoptosis in DMD-derived iPSC cardiomyocytes. Complementary to the phenotype of DMD iPSC-derived cardiomyocytes, DMD patients develop cardiomyopathy, fibrosis and cardiac arrhythmias. Taken together, this corroborates the inventors' findings that deletion of exons 8 and 9 of the DMD gene in normal iPSCs generates an iDMD model, and that cardiomyocytes derived from ΔΕχ8-9 iDMD serve as a model of "DMD in a dish".

The approach reported here is to use CRISPR/Cas9-mediated genomic editing to restore the open reading of the DMD gene and measure the functional outcome of truncated dystrophin in muscle. One of the issues of using CRISPR/Cas9 editing is the occurrence of unintended, off-target, genomic cleavage. Reassuringly, previous reports showed that CRISPR/Cas9- modified human iPSC clones do not exhibit elevated off-target mutation rates, supporting the use of edited iPSC clones for disease modeling. Nevertheless, the inventors recognize that Cas9 nuclease poses a potential safety concern for clinical applicability and acknowledge that whole- genome sequencing should be performed in future in vivo studies to identify potential off-target changes in the genome.

Mutations in the ABD-1 region are genotypically and phenotypically variable in patients. In some cases, out-of-frame ABD-1 mutations display a BMD phenotype, most likely due to the presence of an alternative translation initiation site in exons 6 or exon 8. Interestingly, it was shown that deletion of exon 2 activates an alternative translation initiation site in exon 6 to generate truncated dystrophin. However, duplication of exon 2, which results in a stop codon within the duplicated exon 2, does not activate an alternative translation initiation site. To correct the duplication of exon 2, a U7 small nuclear RNA was used against exon 2 to skip one or both copies of exon 2, generating either full length or truncated dystrophin. Another approach used to correct exon 2 duplication was to use CRISPR/Cas9-mediated editing with one gRNA directed against a duplicated intronic region, resulting in precise deletion of one of the duplicated exons and restoration the DMD gene. Patients with deletion of exons 3-7 display variable phenotypes, depending on whether the alternative translation initiation site in exon 8 is used. However, in general, patients with ABD-1 mutations display a severe BMD phenotype or are diagnosed as DMD. Studies using micro- and mini-dystrophins as therapeutic approaches to treat DMD showed that the truncated dystrophins remain functional if they lack portions of the central rod domain. However, restoration of muscle function by expression of micro- dystrophin in the mdx mouse model of DMD must include an intact ABD-1 region.

Correcting the ABD-1 mutation in DMD is not exclusively based on restoring the open reading frame. Functional assessment of the truncated dystrophin isoforms that are modified in the ABD-1 region is essential. There are three actin-binding sites (ABSs) localized within the ABD-1 region: ABS1 (amino acids 18-27 encoded by exon 2); ABS2 (amino acids 88-116 encoded by exon 5); and ABS3 (amino acids 131-147 encoded by exon 6). The three approaches to correct ΔΕχ8-9 iDMD iPSCs by CRISPR/Cas9-mediated editing produced different modifications to the dystrophin ABD-1 region, such that corrected ΔΕχ3-9 retained the ABS1 ; corrected ΔΕχ6-9 retained both ABS1 and ABS2; and corrected ΔΕχ7-11 retained all three actin binding sites. Unexpectedly, the ΔΕχ7-11 correction, which retained most of the ABD-1 region, created the least stable protein, showing minimal restoration of function. The ΔΕχ7-11 iPSC-derived cardiomyocytes expressed low levels of dystrophin protein compared to isogenic control cells and the corrected ΔΕχ3-9 and ΔΕχ6-9 lines. The decrease in dystrophin expression in the ΔΕχ7-11 iPSC-derived cardiomyocytes resulted in slower Ca ²⁺ cycling and higher propensity for arrhythmias. In eDystrophin (edystrophin.genouest.org), a database dedicated to human dystrophin variants produced by in-frame DMD gene mutations, a patient with the ΔΕχ7-11 DMD deletion was annotated as DMD. It has also been reported that a single missense point mutation in exon 11 causes DMD protein mis-folding and dysfunction, resulting in a BMD phenotype. Therefore, the inventors conclude that although the deletion of exons 7- 11 maintains the open reading frame of the DMD gene, the absence of amino acids 178-444 causes protein mis-folding and subsequent degradation. Indeed, by inhibiting proteasome activity, truncated dystrophin protein increased in ΔΕχ7-11 iPSC-derived cardiomyocytes. Furthermore, it was reported that when recreated in vitro, point mutations in the ABD-1 region that cause a human DMD phenotype are associated with decreased dystrophin protein levels due to proteasomal degradation.

Correction of DMD ΔΕχ8-9 mutation by deleting exons 6-9 retained ABS1 and ABS2 and improved Ca ²⁺ activity in cardiomyocytes, but did not fully restore function to control levels. Shorter time to peak in these cells indicates faster Ca ²⁺ release from the intracellular stores. However, slower Ca ²⁺ decay in these cells compared to control cells suggests that their Ca ²⁺ re-uptake system was not completely restored, resulting in arrhythmias. A similar strategy was applied to correct the ABD-1 mutation in a golden retriever dog model of DMD, which harbors a mutation in the exon 7 splice site. Some functional improvement was seen when exons 6-8 were deleted, resulting in splicing of exon 5 to 9 or to exon 10 due to occurrence of a cryptic splice site in exon 9.

Among the three genome-editing correction strategies the inventors tested, ΔΕχ3-9 which generated the truncated dystrophin lacking amino acid residues 32-320, was the most effective in restoring functionality of iPSC-derived cardiomyocytes. The inventors used CRISPR/Cas9 to create ΔΕχ3-9 using ΔΕχ8-9 iDMD or DMD patient-derived ρΔΕχ3-7 iPSCs, and they restored iPSC-derived cardiomyocyte function in both ΔΕχ3-9 corrected cell lines. ΔΕχ3-9 produces truncated dystrophin that retains ABS1. Consistent with the inventors' findings, it was shown that overexpressing the dystrophin isoform lacking ABS2 and ABS3, prevents severe dystrophy in mdx mice. In addition, by surveying different databases, one group identified 15 patients with an exon 3-9 deletion, 11 of whom were asymptomatic or diagnosed as BMD. In fact, one of these patients is a competitive badminton player and was asymptomatic until he was diagnosed with BMD at age 67. His dystrophin level was recorded using two different dystrophin antibodies, as 47% of the control level using a C-terminus antibody and 62% of the control level using a rod domain antibody. Another of these patients expressed only 15% of normal dystrophin levels and showed a slight decrease in cardiac function at age 21 with no obvious skeletal muscle involvement. Similarly, in mouse models, at least 20% of full-length or central rod domain-deleted dystrophin expression is required to rescue the mdx phenotype. In human mutations involving the 5 '-proximal region of the DMD gene, 30% or more of dystrophin expression is required to prevent diagnosis of muscular dystrophy in X-linked dilated cardiomyopathy patients. Taken together, these reports show in both mice and humans that deletion of exons 3-9 of the DMD gene produces a stable, albeit truncated dystrophin protein capable of maintaining function even when expressed at low levels.

The functional relevance of the ΔΕχ3-9 mutation and its potential therapeutic superiority compared to ΔΕχ6-9 and ΔΕχ7-11 was further underscored by phenotypic screening in EHM. All three models resulted in improved contractile performance compared to the iDMD model, however, ΔΕχ3-9-ΕΗΜ showed the most enhanced contractile force and lowest propensity for arrhythmic contractions. This suggests deletion of exons 3-9 of the DMD gene as a practical candidate for exon-deletion which could be applicable to -7% of DMD patients' mutations. The other two correction strategies that deleted exons 6-9 or 7-11, each could potentially correct -2% of DMD patients' mutations based on the UMD-DMD database. Although shown as a possible gene therapy approach for correcting DMD, the inventors are aware that deletion of exons 3-9 of the DMD gene generates a large genome deletion up to -150 kb. To this point, large genome deletions by CRISPR/Cas9-mediated editing of up to 725 kb of the DMD gene have been previously reported to delete exons 45-55 in human cells in vitro, and in a humanized DMD mouse model in vivo.

To test efficiency of CRISPR/Cas9-mediated deletion of exons 3-9 of the DMD gene by two gRNAs and the function of the truncated dystrophin in vivo, the inventors generated the ΔΕχ8-9 DMD mouse model, which presents dystrophic phenotype. This model could be used to test multiple exon skipping as well as exons refraining approaches to correct dystrophin ABD-1 mutations.

In summary, the inventors' human iPSC results provide evidence that deletion of exons 3-9 in the DMD gene restores muscle function and is applicable for using CRIPSR/Cas9 genomic editing to correct ABD-1 mutations that were previously not addressed. The functional properties of these dystrophin ABD-1 mutations will dictate the most efficacious CRISPR/Cas9 strategy for possible genomic editing to correct DMD mutations within the proximal hot spot of the DMD gene allowing effective restoration of dystrophin function.

VI. Examples

The following examples are included to demonstrate preferred embodiments of the disclosure. It should be appreciated by those of skill in the art that the techniques disclosed in the examples which follow represent techniques discovered by the inventor to function well in the practice of the disclosure, and thus can be considered to constitute preferred modes for its practice. However, those of skill in the art should, in light of the present disclosure, appreciate that many changes can be made in the specific embodiments which are disclosed and still obtain a like or similar result without departing from the spirit and scope of the disclosure.

EXAMPLE 1 - Materials and Methods

Study Approval. All experimental procedures involving animals in this study were reviewed and approved by the University of Texas Southwestern Medical Center's Institutional Animal Care and Use Committee.

Human iPSC maintenance, nucleofection and differentiation. Human healthy donor induced pluripotent stem cells were reprogrammed from peripheral blood mononuclear cells with CytoTune-iPS Sendai Reprograming Kit (catalog # A16518; Thermo Fisher Scientific). Human Muscular Dystrophy iPS Cell Line (catalog #SC604A-MD; Systems Biosciences Inc.) were referred to as ρΔΕχ3-7. Human iPSCs were cultured in mTeSR™l media (catalog #05850; STEMCELL Technologies) and passaged approximately every 3-4 days (1 : 12-1 : 18 split ratio). One hour before nucleofection, iPSCs were treated with 10 μΜ ROCK inhibitor, Y-27632 (catalog #S1049, Selleckchem) and dissociated into single cells using Accutase (catalog #A6964 Innovative Cell Technologies, Inc.). 1 x 10 ⁶ iPS cells were mixed with 6 μg total of pSpCas9(BB)-2A-GFP (PX458) from Feng Zhang (MIT, Cambridge, MA) Addgene plasmid #48138 (Ran et al, 2013) plasmid and nucleofected using the P3 Primary Cell 4D- Nucleofector X kit (catalog #V4XP-3024; Lonza) according to manufacturer's protocol. After nucleofection, iPSCs were cultured in mTeSR™l media supplemented with 10 μΜ ROCK inhibitor and 100 μg/ml Primocin (InvivoGen), and the next day the media was switched to fresh mTeSR™l. Three days post-nucleofection, GFP(+) and GFP(-) cells were sorted by FACS and subjected to genotyping by PCR. Single clones derived from GFP(+) iPSCs were picked, genotyped and sequenced. iPSCs were induced to differentiate into cardiomyocytes, using previously described protocol with modifications. When cells reached -80% confluency the medium was changed to CDM3 every other day until day 10. On day 0-2, medium was supplemented with 4 μΜ of CHIR99021 (catalog #S2924; Selleckchem). On day 2-4 medium was supplemented with 2 μΜ WNT-C59 (catalog #S7037; Selleckchem). On day 10 after differentiation initiation, media was changed to RPMI 1640 medium, no glucose (catalog #11879020; Thermo Fisher Scientific) supplemented with B27-supplement (catalog #17504044; Thermo Fisher Scientific) for 10 days. On day 20 media was switched to cells maintaining media, RPMI 1640 medium, (catalog #11875093; Thermo Fisher Scientific) supplemented with B27-supplement. Cardiomyocytes were used for experiments on day 30-40 after initiation of differentiation.

Genomic DNA isolation. Genomic DNA of human iPSCs was isolated using

DirectPCR (Cell) (catalog #302-C; Viagen) according to manufacturer's recommendations.

RT-PCR. RNA from iPSC-derived cardiomyocytes was isolated using TRIzol (catalog #15596026; ThermoFisher Scientific), and extracted with Direct-zol ^tm RNA MiniPrep kit according to manufacturer's protocol. cDNA was synthesized using iScript ⁰¹¹ gDNA Clear cDNA Synthesis Kit (catalog #1725034; Bio-Rad Laboratories) according to manufacturer's instructions. 2μg of cDNA per reaction was used for RT-PCR reaction using Taq polymerase for 40 cycles. Primer pairs used for human DMD RT-PCR were as follows:

Exon 1 forward: 5 ' -CTTTGGTGGGAAGAAGTAGAGGACTG-3 ' (SEQ ID NO: 83) 5 '-UTR forward: 5'-CTTTCCCCCTACAGGACTCAG-3' (SEQ ID NO: 84)

Exon 5 forward: 5'-TTGGAAGTACTGACATCGTAGATGGA-3' (SEQ ID NO: 85) Exon 10 reverse: 5 ' -CTC AGC AGAAAGAAGCC ACGATAATA-3 ' (SEQ ID NO: 86) Exon 12 reverse" 5 ' -TGTTAGCC AGTC ATTC AACTCTTTC A-3 ' (SEQ ID NO: 87) PCR products for exon 1 forward exon 10 reverse were 1073 bps in Ctrl iPSC-CM 762 bps in AEx8-9 iDMD iPSC-CM and 206 bps in AEx3-9 iPSC-CM. PCR products for exon 5 forward exon 10 reverse were 797 bps in Ctrl iPSC-CM 486 bps in AEx8-9 iDMD iPSC-CM and 194 bps in AEx6-9 iPSC-CM. PCR products for exon 5 forward exon 12 reverse were 1115 bps in Ctrl iPSC-CM 804 bps in AEx8-9 iDMD iPSC-CM and 314 bps in AEx7-l l iPSC-CM. PCR products for 5'-UTR forward exon 10 reverse were 1345 bps in Ctrl iPSC-CM 748 bps in pAEx3-7 iPSC-CM and 544 bps in pAEx3-9 iPSC-CM. PCR products were run on 1.5% agarose gel, excised and sequenced. Primer pair used for human a-actinin control is:

ACTN2 forward: 5 ' -C AACTTC AAC ACGCTGC AGACC AA-3 ' (SEQ ID NO: 88) ACTN2 reverse: 5 ' -AAGCGCTCC AGTCTTCGAATCTC A-3 ' (SEQ ID NO: 89) Dystrophin Western blot analysis. iPSC-CMs were collected in cold PBS on ice, centrifuged and lysed with RIPA lysis buffer: 150mM NaCl, 0.5% sodium deoxycholate, 0.1% SDS, 5% glycerol, 50mM Tris-HCl pH 7.4, 1% Nonidet P40. Samples were needle- homogenized, big clumps were removed by centrifugation and supernatant was incubated at 95 °C 10 min in the presence of Laemmli sample buffer. 10 μg of protein per sample was separated on Mini-PROTEAN TGX 4-20% pre-cast gel (BioRad) for 3 hr at 100V. Proteins were transferred to PVDF membrane at 100V for 1.5 hr. Membranes were probed with mouse anti-dystrophin antibody MANDYS8 (1 : 1000) (catalog #D8168; Sigma) and with mouse anti- vinculin antibody, hVIN-1 (1: 1000) (catalog #V9131; Sigma). For analysis of DEx7-l l iPSC- derived cardiomyocytes treated with proteasome inhibitor MG132, membranes were cut and the upper portion was probed with mouse MANDYS8 anti-dystrophin antibody (1 : 1000) (catalog # D8168; Sigma) and the lower portion with mouse anti-GAPDH antibody, (1: 1000) (catalog # MAB374; 6C5, EMD Milipore). Goat anti-mouse HRP-conjugated secondary antibodies were used (1: 10,000) (catalog #172-1011; Bio-Rad).

Dystrophin and troponin-I immunocytochemistry. iPSC-derived cardiomyocytes (1 x 10 ⁵ ) seeded on 12 mm coverslips coated with poly-D-lysine and Matrigel (catalog #354248; Corning,) were removed from culture media and fixed in cold acetone (10 minutes, -20 °C). Following fixation, coverslips were equilibrated in phosphate-buffered saline, pH 7.3 (PBS) and then blocked for one-hour with serum cocktail (2% normal horse serum/2% normal donkey serum/0.2% bovine serum albumin (BSA/PBS)). Excess blocking cocktail was removed and dystrophin/troponin primary antibody cocktail, mouse anti-dystrophin (1 :800) (catalog # D8168; MANDYS8, Sigma- Aldrich) and rabbit anti-troponin-I (1 :200) (catalog #sc-15368; HI 70, Santa Cruz Biotechnology) in 0.2% BSA/PBS was applied without an intervening wash. Following overnight incubation at 4 °C, unbound primary antibodies were removed with PBS washes, and coverslips were probed for one-hour with secondary antibodies, biotinylated horse anti-Mouse IgG (1 :200) (Vector Laboratories) and fluorescein-conjugated donkey anti-rabbit IgG (1 :50) (Jackson Immunoresearch) diluted in 0.2% BSA/PBS. Unbound secondary antibodies were removed with PBS washes and final dystrophin labeling done with 10-minute incubation of rhodamine-avidin-DCS (1 :60) (Vector Laboratories) diluted in PBS. Unbound rhodamine was removed with PBS washes, nuclei were counterstained with 2 μg/ml Hoechst 33342 (Molecular Probes, Eugene, OR), and coverslips were inverted onto slides overlaying Vectashield fluorescent mounting media (Vector Laboratories).

Calcium imaging. iPSC-derived cardiomyocytes were dissociated with TryplE (catalog #12605036; Thermo Fisher Scientific) and single cells were plated at low density on Matrigel coated 35 mm glass-bottom dishes and Ca ²⁺ handling was measured 3-4 days after plating. To monitor changes in cytoplasmic Ca ²⁺ , iPSC-derived cardiomyocytes were loaded with 5 μΜ fluorescent Ca ²⁺ indicator Fluo-4 AM (catalog # F14201 ; Thermo Fisher Scientific) in the presence of 2 mmol/L of Pluronic F-127 for 20 min in Tyrode's solution (140 mM NaCl, 5.4 mM KC1, 1 mM MgCh, 10 mM glucose, 1.8 mM CaCh, 10 mM HEPES, pH 7.4). The Ca ²⁺ transients of spontaneously beating iPSC-derived cardiomyocytes were imaged at 37 °C using a Zeiss LSM880 confocal system using 40x oil immersion objective. Ca ²⁺ transients were processed and analyzed using ImageJ (NIH, Bethesda) and pClamp 10.2 software (Axon Instrument). For the calcium release phase, the inventors calculated time to peak (time from baseline to maximal point of the transient). For evaluation of calcium re-uptake, the total transient time and time of Ca ²⁺ decay were evaluated. The time of Ca ²⁺ decay was calculated by fitting a first-order exponential decay to the calcium reuptake phase of the calcium transient profile. Cells were considered arrhythmic if they had one or more asynchronized Ca ²⁺ transient. For each experimental group, the inventors analyzed cells from at least three independent sets of iPSC-derived cardiomyocytes.

Engineered heart muscle. Engineered heart muscle (EHM) was constructed as previously reported from iPSC-derived cardiomyocytes of iDMD (ΔΕχ8-9; n=6) and the corrected lines (ΔΕχ3-9, n=14; ΔΕχ6-9, n=3; ΔΕχ7-11, n=3). All lines were adapted to a published cardiomyocyte differentiation protocol, which resulted in high and homogeneous cardiomyocyte purity in the different iPSC cultures, showing 95 ± 1% a-actinin positive (ACTN2 ⁺ ) cardiomyocytes by flow cytometry; n=8. For EHM construction, 1 x 10 ⁶ iPSC- derived cardiomyocytes were mixed with 0.5 x 10 ⁶ human foreskin fibroblasts (catalog #SCRC-1041 ; ATCC). EHM was cultured for 4 weeks followed by isometric force measurements under electrical field stimulation (1.5 Hz) at 37 °C in Tyrode's solution. Two healthy donor-derived iPSC lines were included as a reference: control (WT) shiPS9-l (Daniel Garry, University of Minnesota, Minneapolis, MN) (n=4) and hiPSC-Gl line (n=4).

CRISPR/Cas9-mediated exon deletion in mice. Two single-guide RNA (sgRNA) specific regions of introns 7 and 9, surrounding exons 8 and 9 in the sequence of the mouse Dmd locus, were cloned into vector pX458. For the in vitro transcription of sgRNA, T7 promoter sequence was added to the sgRNA template by PCR. The gel purified PCR products were used as template for in vitro transcription using the MEGAshortscript T7 Kit (Life Technologies). sgRNA were purified by MEGAclear kit (Life Technologies) and eluted with nucl ease-free water (Ambion). The concentration of guide RNA was measured by aNanoDrop instrument (Thermo Scientific).

Genotyping of ΔΕχ8-9 DMD Mice. ΔΕχ8-9 DMD mice were genotyped using primers encompassing the targeted region as follows:

mDMD_i7_forward: 5'-GAGGTTTAAAACATTAAGCCTTTCC-3' (SEQ ID NO: 90) mDMD_i7_reverse: 5 ' - AC ATTAAGATGGACTTCTTGTCTGG-3 ' (SEQ ID NO: 91) mDMD_i9_reverse: 5 ' -TACTC ACATATGGGTCGTTTTCCTT-3 ' (SEQ ID NO: 92).

Tail biopsies were digested in 100 of 25 mM NaOH, 0.2 mM EDTA (pH 12) for 20 min at 95 °C. Tails were briefly centrifuged followed by addition of 100 of 40 mM Tris HCl (pH 5) and mixed to homogenize. Two microliters of this reaction was used for subsequent PCR reactions with the primers below, followed by gel electrophoresis.

Histological analysis of muscles. Skeletal muscles from WT and ΔΕχ8-9 DMD mice were individually dissected and cryoembedded in a 1 :2 volume mixture of Gum Tragacanth powder (Sigma- Aldrich) to Tissue Freezing Medium (TFM) (Triangle Bioscience). All embeds were snap frozen in isopentane heat extractant supercooled to -155 °C. Resulting blocks were stored overnight at -80 °C prior to sectioning. Eight-micron transverse sections of skeletal muscle, and frontal sections of heart were prepared on a Leica CM3050 cryostat and air-dried prior to same day staining. H&E staining was performed according to established staining protocols and dystrophin immunohistochemistry was performed using MANDYS8 monoclonal antibody (Sigma-Aldrich) with modifications to manufacturer's instructions. In brief, cryostat sections were thawed and rehydrated/delipidated in 1% triton/phosphate-buffered-saline, pH 7.4 (PBS). Following delipidation, sections were washed free of Triton, incubated with mouse IgG blocking reagent (M.O.M. Kit, Vector Laboratories), washed, and sequentially equilibrated with MOM protein concentrate/PBS, and MANDYS8 diluted 1 : 1800 in MOM protein concentrate/PBS. Following overnight primary antibody incubation at 4 °C, sections were washed, incubated with MOM biotinylated anti-mouse IgG, washed, and detection completed with incubation of Vector fluorescein-avidin DCS. Nuclei were counterstained with propidium iodide (Molecular Probes) prior to cover slipping with Vectashield.

Statistical analysis. Statistical analysis was assessed by one-way ANOVA or two-way ANOVA and Tukey's post-hoc test (F) were indicated. Data are shown as mean ± SEM. A PO.05 value was considered statistically significant.

EXAMPLE 2 - Results

Generating a dystrophin mutation that disrupts the actin-binding domain. The inventors generated a human iPSC line with a deletion of exons 8 and 9 in the DMD gene using CRISPR/Cas9-mediated editing to analyze the effect ABD-1 deletions on muscle function (FIG. 1A). Two single guide RNAs (gRNAs), targeting sequences in intron 7 and intron 9 of the DMD gene in the presence of Streptococcus pyogenes Cas9, were used to delete DMD exons 8-9 in healthy human iPSCs (FIG. IB). This induced DMD mutant human cell line is referred to as ΔΕχ8-9 iDMD, and represents the common 5 '-proximal hot spot mutations. Human iPSCs harboring DMD exons 8-9 deletions were picked by clonal selection and the mutation was confirmed by sequencing genomic DNA (FIG. 1C). The human ΔΕχ8-9 iDMD iPSC line has a 1,909 bp deletion, which generates a newly formed junction between intron 7 to intron 9 (FIG. 1C). Primers within intron 7 flanking the gRNA-7 targeting site were used to validate the deletion and as expected showed no PCR product with ΔΕχ8-9 iDMD compared to control iPSC with a 424 bp product (FIG. 1C). The AEx8-9 iDMD iPSCs were differentiated to cardiomyocytes and RT-PCR was performed using forward and reverse primers targeting exons 1 and 10, respectively (FIG. ID). Sequencing of the RT-PCR products showed splicing of exon 7 to exon 10, which created a premature stop codon following the first 5 amino acids encoded by exon 10 (FIG. ID). Loss of dystrophin protein was confirmed by Western blot analysis and immunocytochemistry in ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes (FIGS. 1E-F).

Correction of the AEx8-9 iDMD mutation by exon deletion strategies. Three different CRISPR/Cas9-mediated strategies were used to correct the mutation in ΔΕχ8-9 iDMD iPSCs in order to restore the dystrophin open reading frame (FIG. 2A). These strategies generated dystrophin with different truncations in the ABD-1 domain, providing a means of analyzing the effect of ABD-1 deletions on muscle function. For each strategy, the inventors applied two gRNAs to delete multiple exons and analyzed two independent iPSC clones. Strategy #1 generated ΔΕχ3-9 iPSCs, by targeting intron 2 and intron 7 with gRNAs, and deleting exons 3 through 7 (FIG. 2B). Genomic sequencing of the genome edited region showed a hybrid junction of intron 2 to intron 7, confirming generation of ΔΕχ3-9 iPSCs (FIG. 2C). Following differentiation of ΔΕχ3-9 iPSCs to cardiomyocytes, RT-PCR was performed using primers within exon 1 and exon 10 (FIG. 2D). Sequencing of the RT-PCR product showed splicing of exon 2 to exon 10. Restoration of the DMD open reading frame generated a truncated dystrophin protein lacking the ABS-2 and ABS-3 regions. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry of ΔΕχ3-9 iPSC-derived cardiomyocytes (FIGS. 2E-F; FIG. 7A).

Strategy #2 generated ΔΕχ6-9 iPSCs, using gRNAs targeting intron 5 and intron 7 in order to delete exons 6 and 7 (FIG. 3 A). This form of CRISPR/Cas9 editing resulted in deletion of exon 6 through exon 9, which was confirmed by sequencing of genomic PCR products (FIG. 3B). Following differentiation, RT-PCR using primers within exon 1 and exon 10 was performed on ΔΕχ6-9 iPSC-derived cardiomyocytes and sequencing of the RT-PCR product confirmed splicing of exon 5 to exon 10 (FIG. 3C). Restoration of dystrophin expression generated a truncated protein lacking the ABS-3 region. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry in ΔΕχ6-9 iPSC-derived cardiomyocytes (FIGS. 3D-E; FIGS. 7A-B).

Strategy #3 generated ΔΕχ7-11 iPSCs, using gRNAs targeting introns 6 and 11 to delete exons 7 to 11 of ΔΕχ8-9 iDMD iPSCs (FIG. 4A). This genome editing strategy introduced the largest deletion of 164 kb in ΔΕχ8-9 iDMD iPSCs. Similar to the first two correction strategies, single cell-derived colonies were picked from the corrected iPSC pool and the genomic region was sequenced to confirm editing (FIG. 4B). The ΔΕχ7-11 iPSCs were differentiated to cardiomyocytes, and RT PCR was performed with primers within exon 5 and exon 12 to reveal splicing of exon 6 to exon 12, as seen by sequencing (FIG. 4C). Restoration of the DMD open reading frame of ΔΕχ7-11 iPSC-derived cardiomyocytes generated a truncated dystrophin protein lacking amino acids 177-444 but with all three ABS intact. Dystrophin expression was confirmed by Western blot analysis and immunocytochemistry (FIGS. 4D-E; FIGS. 7A-B). The inventors observed less dystrophin expression in ΔΕχ7-11 iPSC-derived cardiomyocytes compared to control iPSC-derived cardiomyocytes and the other corrected lines, ΔΕχ3-9 and ΔΕχ6-9. Previously, it was shown that missense mutations associated with the DMD phenotype caused steady-state decreases in dystrophin protein levels inversely proportional to the tertiary stability and directly caused by proteasomal degradation. Consistent with this report, the inventors observed increased dystrophin protein levels when ΔΕχ7-11 iPSC-derived cardiomyocytes were treated with the proteasome inhibitor MG-132, as measured by Western blot analysis (FIG. 4F; FIG. 7C). This suggests that deletion of DMD exons 7-11, which produces dystrophin lacking 177-444 amino acids, results in a truncated form of dystrophin that is degraded.

Functional restoration of iPSC-derived cardiomyocytes by correction of ABD-1 mutations. To assess the consequences of ABD-1 mutations and the effect of correction by genome editing on dystrophin function in muscle, the inventors analyzed spontaneous Ca ²⁺ activity in iPSC-derived cardiomyocytes (FIG. 5A). For each CRISPR/Cas9-edited correction, the inventors analyzed data sets from two corrected clones. As expected, the inventors observed that calcium release and reuptake parameters, including time to peak, Ca ²⁺ decay rate, and transient duration were significantly higher in the ΔΕχ8-9 iDMD iPSC-derived cardiomyocytes compared to isogenic control cells (FIGS. 5B-D). Additionally, greater than 50% of the ΔΕχ8- 9 iDMD iPSC-derived cardiomyocytes demonstrated abnormal Ca ²⁺ activity, which represents an arrhythmogenic-like phenotype (FIG. 5E). In contrast, only 10.7% of the control iPSC- derived cardiomyocytes were desynchronized (FIG. 5E).

Corrected ΔΕχ3-9 iPSC-derived cardiomyocytes displayed normalized Ca ²⁺ transient kinetics and decreased asynchronous activity similar to isogenic control cells, suggesting restoration of dystrophin function (FIGS. 5A-G). Cardiomyocytes derived from ΔΕχ6-9 iPSCs showed improved Ca ²⁺ handling, although not to the level seen with ΔΕχ3-9 cells (FIGS. 5A- G). The Ca ²⁺ decay rate and Ca ²⁺ transient duration were shorter in ΔΕχ6-9 iPSC-derived cardiomyocytes (FIGS. 5B and 5D) and the number of arrhythmic cells was decreased to 29% compared to ΔΕχ8-9 iDMD cells (FIG. 5E). However, the parameters observed in the ΔΕχ6-9 iPSC-derived cardiomyocytes did not reach the levels of control iPSC-derived cardiomyocytes. The ΔΕχ7-11 iPSC-derived cardiomyocytes, similarly to ΔΕχ6-9 cells, showed improvement in Ca ²⁺ release and uptake when compared to ΔΕχ8-9 iDMD cardiomyocytes (FIGS. 5B and 5C), however they did not reach control cell levels. iPSC-derived cardiomyocytes with ΔΕχ7- 11 had the highest number of arrhythmic cells, up to 35.9%, among the corrected cell lines (FIG. 5E). Taken together, these findings indicate that deletion of exons 3-9 (ΔΕχ3-9) to correct DMD mutations within the 5 '-proximal hot spot of the DMD gene produces a truncated dystrophin protein with restoration of function. Further assessment of the functional properties of the iPSC-derived cardiomyocyte cell lines was performed by generating engineered heart muscle (EHM), which displays phenotypic properties of the postnatal myocardium. EHM was generated as shown in FIG. 8A using -95% pure iPSC-derived cardiomyocytes, as measured by a-actinin expression (FIG. 8B). Consistent with the calcium handling data, the inventors observed enhanced contractile performance in all corrected cell lines, with the most pronounced effects in ΔΕχ3-9 EHM (FIG. 5F). Moreover, the inventors observed an increased tendency for arrhythmic contractions in ΔΕχ8-9 iDMD- EHM and a reduction in ΔΕχ3-9 EHM (FIG. 8C).

Correction of DMD patient-derived iPSCs by deleting exons 3-9. An iPSC line, referred to as ρΔΕχ3-7, was generated from a DMD patient (SC604A-MD) with a deletion of exons 3-7 in the DMD gene. ρΔΕχ3-7 iPSCs are used as patient-in-a-dish model of DMD with a mutation in the ABD-1. To evaluate the effectiveness of exon 3-9 deletion in restoration of cardiomyocyte function, the inventors corrected ρΔΕχ3-7 by deleting exons 8 and 9 to generate ρΔΕχ3-9. Two gRNAs targeting introns 7 and 9 were used to excise exons 8-9, generating ρΔΕχ3-9 iPSCs (FIG. 6A). Sequencing of genomic DNA PCR products confirmed the hybrid junction of intron 7 to intron 9 in a single cell-derived iPSC colony (FIG. 6B). Both ρΔΕχ3-7 and ρΔΕχ3-9 iPSC lines were differentiated into cardiomyocytes and RT-PCR was performed with primers targeting the 5'-UTR and exon 10. In ρΔΕχ3-7 iPSC-derived cardiomyocytes, the junction of exon 2 to exon 8 was seen by sequencing of the RT-PCR product (FIG. 6C). In corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes, the junction of exon 2 to exon 10 was demonstrated by sequencing the RT-PCR product (FIG. 6C). Western blot analysis showed a low level of dystrophin expression in DMD ρΔΕχ3-7 iPSC-derived cardiomyocytes (FIG. 6D). This is consistent with previous reports showing low levels of dystrophin protein expression in patients lacking exons 3-7 due to re-initiation of translation at an in-frame start codon in exon 8 (Fletcher et al, 2012; Winnard et al, 1995). Corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes showed dystrophin expression similar to normal control cardiomyocytes (FIG. 6D). These findings were confirmed by immunocytochemistry (FIG. 6E).

Spontaneous Ca ²⁺ activity was assessed in these patient-derived cells as a measure of iPSC-derived cardiomyocyte functionality (FIGS. 6F-I). In ρΔΕχ3-7 DMD iPSC-derived cardiomyocytes, Ca ²⁺ time to peak and time to half decay were elevated (FIGS. 6F-G). This caused an overall slower Ca ²⁺ transient (FIG. 6H) and elevated the number of arrhythmic cells, up to 53% (FIG. 61). Corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes had significantly improved time to peak and faster Ca ²⁺ decay (FIGS. 6F-G). Although these parameters of the corrected ρΔΕχ3-9 iPSC-derived cardiomyocytes did not reach normal control levels, the number of arrhythmic cells was 18.5%, similar to control iPSC-derived cardiomyocytes (FIG. 61). These findings show improved iPSC-derived cardiomyocyte function after genomic editing to excise exons 3-9 of the DMD gene in DMD patient-derived iPSCs.

ΔΕχ8-9 DMD mouse models recapitulate muscle dystrophy phenotype. To investigate CRISPR/Cas9-mediated exon skipping and refraining in vivo, a mimic of the human ABD-1 region mutations was generated in a mouse model by deleting exons 8 and 9, using CRISPR/Cas9 system directed by two single guide RNAs (sgRNA) (FIG. 9A and Table 2). The inventors designed and validated sgRNAs targeting introns, flanking 5' end and 3' ends of Dmd exons 8 and 9 respectively. C57BL/6 zygotes were co-injected with in vitro transcribed Cas9 mRNA and in vitro transcribed sgRNAs, and then re-implanted into pseudo-pregnant females.

The deletion of Dmd exons 8-9 was confirmed by DNA genotyping. Mice lacking exons 8-9 showed pronounced dystrophic muscle changes in 1 and 2 month-old mice (FIG. 9B). The deletion of these exons placed the dystrophin gene out of frame leading to the absence of dystrophin protein in skeletal muscle and heart (FIG. 9C). The grip strength of the ΔΕχ8-9 DMD was significantly weaker than that in wild-type mice (FIG. 9D). Serum analysis of the ΔΕχ8-9 DMD mice shows a significant increase of creatine kinase (CK) level, which is a sign of muscle damage (FIG. 9E). Taken together, dystrophin protein expression, muscle histology, muscle strength and serum creatine kinase level validated the dystrophic phenotype of the ΔΕχ8-9 DMD mouse model.

All of the compositions and/or methods disclosed and claimed herein can be made and executed without undue experimentation in light of the present disclosure. While the compositions and methods of this disclosure have been described in terms of preferred embodiments, it will be apparent to those of skill in the art that variations may be applied to the compositions and/or methods and in the steps or in the sequence of steps of the method described herein without departing from the concept, spirit and scope of the disclosure. More specifically, it will be apparent that certain agents which are both chemically and physiologically related may be substituted for the agents described herein while the same or similar results would be achieved. All such similar substitutes and modifications apparent to those skilled in the art are deemed to be within the spirit, scope and concept of the disclosure as defined by the appended claims. VII. References

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