Submitted for Review: Sep 29 2015 Published: Dec 10 2015

Duchenne Muscular Dystrophy (DMD) is an hereditary, X-linked neuromuscular disease, resulting from mutations across the DMD gene. The subsequent absence of dystrophin protein prevents the correct formation of the dystrophin-associated protein complex (DAPC), a structural link between the intracellular actin and extracellular matrix. This compromises muscle stability and contractility, giving rise to progressive muscle wasting, the prominent clinical feature of this disease. Over time, muscle deterioration results in loss of ambulation, decline in respiratory and cardiac function and ultimately premature death between the second and third decade of life. Whilst the genetic basis of the disease is well established, care remains palliative and there is a clear unmet medical requirement for a gene therapy to address the underlying cause of this devastating disease [1]. A number of different gene therapies are currently in clinical trials and are reporting varying degrees of therapeutic benefit (reviewed in [2,3]). Briefly, these include adeno-associated virus (AAV) microdystrophin delivery [4], premature termination codon read-through using ataluren (Translarna^TM) [5], exon-skipping [6–8] and utrophin upregulation [9,10]. These gene therapies would require repeat administration, and/or carry an adverse immunological risk, and/or are restricted by mutation specificity, all of which may limit their clinical relevance. Such problems may be circumvented through the use of genome engineering.

Therapeutic engineering of the DMD gene

Although the potential benefit of genome engineering strategies to correct the DMD gene is immense, this approach is challenged by the large size of the gene and the diversity of genetic mutations found in patients [11]. The mutation subsets and their relative occurrences are: large deletions 68%, duplications involving more than one exon 12%, and small mutations 20%, as reported by the Treat-NMD Global database [12]. These mutations do not occur uniformly across the length of the DMD gene; they are rather clustered into minor and major hotspots, between exons 2–20 and 45–55 respectively [3,13]. Genome editing could be used to affect permanent correction of numerous mutations via relatively few therapeutic strategies utilizing DNA repair mechanisms. This process orchestrates permanent changes in the mutated gene and, once corrected, the gene is expressed at its endogenous levels. Such correction of the primary genetic defect would prevent the necessity of repeated administration associated with traditional gene therapy approaches, and thus the likelihood of an adverse immunological reaction. Moreover, this can be used to target frequently occurring mutations or indeed intragenic mutational hotspots, excluding multiple exons and thus multiple mutations in the local region. The net result would produce a shortened functional dystrophin protein, whilst also increasing the therapeutic applicability of such an approach.

Genome engineering approaches classically depend upon the use of artificially engineered nucleases with the propensity to introduce a double strand break (DSB) at a defined location within the genome. Initially, this requirement was fulfilled by zinc finger nucleases (ZFNs), meganucleases (MGNs) and transcriptional activator-like effector nucleases (TALENs) [14–16]. ZFNs and TALENs became favored platforms, however they require two engineered protein motifs to confer specific binding, upstream and downstream of a DNA locus of interest, and the dimerization of FokI nucleases to cleave the DNA [17]. Although this represented a major advancement within the field, there were inherent limitations pertaining to the design of constructs being time consuming, expensive and resulting in variable efficacy. The subsequent discovery of CRISPR/Cas9 as a gene editing platform completely revolutionized the field due to the simplicity of guide design and its versatility [18,19].

Genome engineering is affected through the repair of the DSB. In the absence of a donor DNA template, repair via non-homologous end joining (NHEJ) will occur. This is an error prone method of DNA repair perpetuating small nucleotide insertions or deletions (INDELs), which has been shown to restore the reading frame and full-length or partial dystrophin protein expression [20,21] (Figure 1 Schematic diagrams for the use of CRISPR/Cas9 to fix a variety of small DMD mutations.A & C). A shortcoming of this approach is that the length of subsequent INDELs introduced at the DSB are variable in nature; thus the restoration of the reading frame and by extension dystrophin occurs by chance and cannot be precisely controlled. Permanent removal of exons mediated by this process may however enable the restoration of DMD reading frame and prove therapeutic. The removal of the mutational hotspot between exons 45–55 and the excision of exon 51, which could have applicability to 62% and 13% of DMD patients respectively, are two important demonstrations of this therapeutic strategy [22,23] (Figure 2B). An analogous method could also be used to remove other mutation hotspots or indeed exon duplications (Figure 1C). In the latter, the cut sites may flank the duplication or indeed be directed to a region common to both endogenous and duplicated intron sequences, so that the subsequent DSB result is a single copy of each exon. In contrast, homology-directed repair (HDR) rectifies a DSB through the addition of genetic material when a homologous repair template is supplied. The simplest DNA donor currently utilized is a single stranded oligonucleotide (ssODN), which has been used by multiple groups to introduce subtle sequence alterations in genes other than DMD [24–26] (Figure 1B). Although ssODNs are often used to establish constitutive levels of HDR, they are being used increasingly to correct point mutations, one prominent example being the correction of the nonsense stop mutation carried by the mdx mouse, the established animal model of DMD [27]. Whilst the level of HDR is known to vary amongst different cell types, this is a promising approach for the permanent correction of small DMD mutations. It should be noted that this approach can only be applied to dividing cells, implicating that it would only be efficient as ex-vivo correction in adult skeletal muscle. However, it could eventually prove to be an alternative to pharmacological read-through reagents such as aminoglycosides and ataluren, which require repeated administration [28,29].

In contrast to NHEJ, HDR using ssODNs could specifically correct the DMD reading frame; insertion of the correct nucleotide(s) at the correct location will allow formation of the correct triplet codon and hence inclusion of the correct amino acid at the required position in the dystrophin protein. The delivery of extended pieces of coding DNA (cDNA) in the form of DNA donor repair templates have been used to address larger deletions. Our group previously demonstrated that a deletion of exons 45–52 was specifically reparable through use of a MGN and designed repair template [30] (Figure 2A) HDR-mediated repair utilizing a cDNA template can be used to insert regions of the DMD gene that are deleted. In this instance, production of a DSB and HDR repair using a donor cDNA template containing exons 45-52 can restore the reading frame and full-length dystrophin expression. B) Multiplexing CRISPR guides can be used to remove exons thereby restoring the reading frame and expresion of a truncated form of dystrophin. The deletion of exons 50–52 in this example can be overcome by cutting out exons between intron 44 and 55. The removal of the exons included within this region enables the restoration of the reading frame, with exons 44 and 56 being in-frame. This removes a mutational hotspot and could be used to treat 62% of DMD patient mutations. DSB: Double-strand break; HDR: Homology-directed repair.A). It should be noted however that the efficiency of HDR is low since it is restricted to the S and G2 phases of the cell cycle [31]. This, together with off-target effects, problems with delivery and guide RNA (gRNA) production, constitute hurdles that need to be overcome before CRISPR/Cas9-mediated genome engineering can realize its potential as a therapy for DMD and other genetic diseases. CRISPR/Cas9 carries a much higher risk of off-target mutations relative to ZFNs and TALENs; these off-target mutations can result in cell death or transformation. To reduce these off-target effects there are four main aspects related to CRISPR/Cas9 that can be further optimized and warrant further research:

Careful selection of target site with consideration of the number of mismatches between gRNA and complementary sequence, their position and distribution is required [32–34].

• gRNAs should be designed to avoid poly-G, poly-C and poly-A rich sequences and where possible truncated gRNAs (17–18 bps) should be used since these have lower rates of off-target effects as a result of decreased mismatch tolerance [24]. gRNA secondary structure, which has the potential to prevent non-target interactions, and also chromatin levels of target should be taken into account [35]. Improved gRNA design tools are consistently being developed and should be exploited. The most recent design tools comparison reports that the sequence preference for CRISPR-mediated transcriptional regulation is substantially different from that for CRISPR/Cas9 knockout [36]. It has become increasingly apparent that Cas9 proteins can be engineered with higher specificity through alteration of their protospacer adjacent motif (PAM) dependence. For example, the recently described Neisseria meningitides Cas9 protein and Streptococcus thermophilus Cas9 proteins require PAMs with sequence 5’-NNNNGATT-3’ and 5’-NNAGAAW-3’, respectively offering much higher stringency in targeting than the established Streptococcus pyrogenes Cas9 (5’-NGG-3’) [18,37,38].
• Transfection conditions can have a dramatic effect on the level of off-target mutations as there is a clear relationship between gRNA and Cas9 concentration and undesired targeting [32,34].
• An examination of the reliability of existing techniques and the development of improved methodology for the detection of off-target mutations.

It should be noted that it has been reported that the use of paired nickases based on dead Cas9 (dCas9) as opposed to single wild-type Cas9 will increase target specificity by 1500 times [39,40], since off-target single nicks are faithfully repaired [41]. The fusion of FokI catalytic domain to dCas9 provides an additional layer of stringency since FokI has rigid spatial requirements for activity and will not cleave as a monomer [42,43].

For delivery, currently DNA (as plasmids expressing Cas9) and gRNA [44] and RNA [45,46] injection-based techniques are used for CRISPR/Cas9 delivery into organisms which is inefficient and non-specific. Efficiency of delivery would be increased through the use of vectors. To allow packaging of AAV vectors which are limited by the size of genetic cargo they can carry, smaller Cas9 orthologs from other microbes have recently been developed [47]. The use of the transfer RNA (tRNA) promoter expression strategy [48] could also greatly facilitate the construction of effective AAV-based Cas9/sgRNA vectors for future in vivo use. The use of AAVs to deliver CRISPR would allow the use of muscle-specific serotypes to enhance Cas9 and gRNA expression in muscle. Other vectors being exploited for CRISPR/Cas9 delivery include adenoviral [49–51] and lentiviral vectors [50,52]. Very recently, two groups have examined the potential of splitting the full wild-type Cas9 coding sequence between two AAV vectors to good effect [53,54]. The development of delivery systems that allow tissue- and cell-specific expression of CRISPR/Cas9 components has recently been demonstrated in zebrafish [55]. It is therefore hoped that muscle-specific expression is potentially possible. Other strategies to enhance delivery of CRISPR/Cas9 include the use of Cas9 recombinant protein [25] and cell-penetrating peptide (CPP) conjugates of this protein together with CPP–gRNA complexes [56]. Chemical alterations to synthesized gRNAs have recently been reported to enhance genome editing efficiency in human primary T-cells and CD34⁺ hematopoietic stem and progenitor cells without the toxicity associated with DNA delivery [57]. A novel delivery vehicle for CRISPR/Cas9 based on a biologically inspired yarn-like DNA nanoclew and synthesized by rolling circle amplification with palindromic sequences encoded to drive the self-assembly of nanoparticles has very recently been described [58]. To allow the precise temporal control of Cas9 gene expression, a light-inducible, user-defined, endogenous gene activation system based on CRISPR/Cas9 has been developed [59]. Application of these recent developments in optimizing delivery, targeting and expression to DMD gene editing should progress the work along the path to clinical readiness.

Strategies to tackle off-target effects, delivery problems as well as improvements in gRNA production and efficiency of HDR for full DMD gene repair are summarized in Table 1. 

Generation of DMD animal models

The availability of emerging CRISPR-based gene editing strategies, with higher specificity and efficiency, has in turn impacted the availability and the development of animal models. In the DMD field, the mdx mouse is still the most widely known and used model since its discovery in the mid-’80s [60]. It carries a nonsense point mutation in exon 23 that causes dystrophin deficiency; however the disease has a milder and non-progressive course compared to the human counterpart. To achieve a closer clinical phenotype than that observed in the mdx mouse, large animals models are required [61]. The CRISPR/Cas9 system has been used for the generation of highly efficient, heritable, gene knockout in mice and rats, where the system has also been used to induce double-gene knockout with only a single microinjection of Cas9 and a mixture of sgRNAs into one-cell-stage rat embryos [62]. Importantly, CRISPR/Cas9 technology is also proven to be effective in non-traditional animal models, like goats and pigs, which are important for both agricultural and biomedical research and development [63–68]. This technology has facilitated simplified genetic modification [69] and creation of disease models [70] in non-human primates. Most recently, Chen and colleagues used CRISPR/Cas9 to target the monkey DMD gene to create mutations in order to recapitulate DMD. In their study, Cas9 induced mosaic mutations in over the 87% of the DMD alleles in muscle, which is also supported by the depletion of dystrophin protein in the targeted muscles and associated muscle pathology [70]. Interestingly, myostatin, a negative regulator of skeletal muscle mass, has also recently been targeted with Cas9/sgRNA by Zou Q et al, to produce a myostatin knock-out dog model [71]. Importantly, concerns arise due to the presence of mosaic mutations produced with the CRISPR system, which might confound the potential phenotype, especially since primates and large animals have much longer breeding times than rodents. Nonetheless, when developing DMD animal models, this aspect could turn out to be useful in acquiring deeper insights into the percentage of dystrophin-positive fibers required to restore the dystrophic phenotype.

This system has also been used for the generation of DMD-mutated rats [72], where the simultaneous targeting of DMD exon 3 and exon 16 resulted in the absence of dystrophin expression in the F0 generation. These mutations were heritable by the next generation, and F1 male rats exhibited similar phenotypes in their skeletal muscles. This demonstrates how two types of mutation apparent in human DMD can be relatively quickly recapitulated producing helpful platforms to explore potential repair strategies. The DMD rat shows a more marked decline in muscle strength, presence of degenerative/regenerative cycles in the heart and diaphragm and more marked muscle fibrosis; it could therefore serve as a superior model in terms of disease pathology to the mdx mouse that is currently so widely used for the testing of therapies for DMD.

The generation of the transgenic humanized mdx mouse model [73] has provided a valuable setting to test the in vivo efficacy of human-specific antisense oligonucleotides (AOs) prior to clinical trial [74–76]. This model expresses full-length human dystrophin protein on an mdx background. Use of an AO to induce exon skipping will disrupt the transcript reading-frame and thereby block dystrophin protein expression. The ideal animal model for testing AO efficacy would be one that is transgenic with human DMD gene carrying a relevant mutation. The use of CRISPR/Cas9 engineering of the humanized mdx mouse to generate such mouse models is yet to be executed but could provide improved translational development of therapies.

Animal models allow a means to investigate the efficacy of both ex vivo and in vivo genome engineering as a therapy for DMD. Yet to be performed, ex vivo genome-engineered stem cells could be engrafted onto dystrophic muscle and amelioration of disease phenotype assessed. In vivo genome correction of the mutation in the germline of the mdx mice using a designed sgRNA to target mdx exon 23 and an ssODN as a template for HDR-mediated gene repair has been reported [27]. The mdx zygotes were co-injected with Cas9 mRNA, sgRNA, and ssODN, and then implanted into pseudo-pregnant female mice. The “corrected” mdx progeny displayed from 2 to 100% correction of the mdx gene. Interestingly, the correction of only 17% of the mutant mdx alleles was sufficient to allow dystrophin expression in a majority of myofibers at a level of intensity comparable to that of wild-type mice, and the muscle exhibited fewer histo-pathologic hallmarks of muscular dystrophy than mdx muscle. Very recently, Xu et al have reported the excision of a 23-kb genomic region of the X-chromosome covering the mdx exon 23 mutation. In contrast to the previous approach described, this was performed using direct in vivo administration of CRISPR/Cas9 into post-natal mdx mice, but similarly resulted in truncated yet functional dystrophin protein being expressed [77].

Altogether these in vivo animal studies have begun to demonstrate a proof of concept that CRISPR/Cas9-based genome editing has the potential for pre-clinical development. Additionally, they demonstrate the potential to target specific tissues and regenerate transient models of disease without the breeding of engineered animals. The application of CRISPR/Cas9 technology for genome editing in a wide range of organisms will promote our understanding of disease pathology and provide animal models with therapeutic relevance for human diseases including DMD.

Unexplored potential of genome engineering for DMD

CRISPR/Cas9-based genome editing does not just hold potential for the correction of the DMD gene, but could also be used to address the multi-factorial nature of the disease using alternative targeting strategies. Cas9 does not necessarily have to be used to cleave DNA; an engineered Cas9 mutant with specific point mutations in its endonuclease domains such that it has no catalytic activity against DNA has been developed [78,79]. Rather than cleaving its target, this so called dCas9, upon binding to its target DNA upstream of a promoter through a gRNA, will prevent RNA polymerase binding and activity, and hence block transcription. It is therefore possible to use Cas9 as a modulator of transcription in a target-specific fashion. Since DMD is a multi-factorial disease in terms of the dystrophic, atrophic, fibrotic skeletal muscle phenotype, this dCas9-mediated transcriptional repression could be exploited. Cas9 can be used to address the dystrophin deficiency through NHEJ/HDR repair of the DMD gene as discussed above, and in its dCas9 mutated form can additionally be used to repress the transcription and hence expression of fibrogenic genes and negative regulators of muscle growth. Such an exciting strategy has not yet been explored with DMD, but could hold great therapeutic potential. It should be noted that the ability to target more than one gene with a single delivery vector has been made possible by the development of lentiviral [52,80] and AAV [81] vector systems for the modular delivery of multiple gRNAs.

Enhancement of gene expression is also possible using dCas9 through its tethering to transcriptional activators, such as VP64, the tetramer of the herpes simplex activation domain VP16 [78,82–84]. As a therapeutic strategy for DMD, VP64-tethered dCas9 could be used to activate the expression of genes involved in myogenesis, and could provide an exciting alternative to small molecule upregulation of utrophin [43], a dystrophin homolog.

For the regulation of expression of fibrogenic/anti-myogenic genes as potential adjunct therapies for DMD using CRISPR/Cas9, the targeting of RNA rather than DNA could hold distinct advantages. It could allow more specific repression/activation and potentially more efficient Cas9 binding. The identification of i) repression of a specific transcript by Francisella novicida Cas9 (FnCas9), guided by a small CRISPR/Cas-associated RNA [85] and ii) RNA targeting by a distinct CRISPR/Cas subtype (Type III) present in Pyrococcus furiosus [86] could provide the basis for programmable Cas9-mediated RNA interference of fibrotic/anti-myogenic transcripts.

With the appropriate gRNA design, CRISPR/Cas9 technology not only holds the potential to permanently modify all DMD mutations, irrespective of type, such that dystrophin protein expression is restored, but also to target associated dystrophic muscle phenotypes to provide an all-encompassing therapy. Application of recent developments in optimizing delivery, targeting and expression to DMD genome engineering should progress the work along the path to clinical readiness. The exponential growth of research resulting from the discovery of CRISPR/Cas9 may in the long term enable the effective cure of monogenic hereditary conditions including DMD, which previously appeared unachievable.

Financial & competing interests disclosure

The authors have no relevant financial involvement with an organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript. This includes employment, consultancies, honoraria, stock options or ownership, expert testimony, grants or patents received or pending, or royalties.
 No writing assistance was utilized in the production of this manuscript.

Acknowledgements

The authors gratefully acknowledge Muscular Dystrophy UK, Duchenne Parent Project and AFM-Telethon for their valued support of the gene editing work performed at Royal Holloway University of London.

This work is licensed under a Creative Commons Attribution- NonCommercial – NoDerivatives 4.0 International License.

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Affiliations

Marc Moore*, Denis Vallese*, George Dickson & Linda Popplewell§
School of Biological Sciences,
Royal Holloway University of London, Egham, Surrey, TW20 0EX, UK
§Author for Correspondence
linda.popplewell@rhul.ac.uk
*Contributed equally to the work.