Advances in CRISPR/Cas9 Genome Editing for the Treatment of Muscular Dystrophies

Abstract

Muscular dystrophies (MDs) comprise a diverse group of inherited disorders characterized by progressive muscle loss and weakness. Given the genetic etiology underlying MDs, researchers have explored the potential of CRISPR/Cas9 genome editing as a therapeutic intervention, resulting in significant advances. Here we review recent progress on the use of CRISPR/Cas9 genome editing as a potential therapy for MDs. Significant strides have been made in this realm, made possible through innovative techniques such as precision genetic editing by modified forms of CRISPR/Cas9. These approaches have shown varying degrees of success in animal models of MD, including Duchenne muscular dystrophy (DMD), congenital muscular dystrophy type 1A (MDC1A), and myotonic dystrophy type 1 (DM1). Even so, there are several challenges facing the development of CRISPR/Cas9-based MD therapies, including the targeting of satellite cells, improved editing efficiency in skeletal and cardiac muscle tissue, delivery vehicle enhancements, and the host immunogenic response. While more work is needed to advance CRISPR/Cas9 genome editing past the preclinical stages, its therapeutic potential for MD is extremely promising and justifies concentrated efforts to move into clinical trials.

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Advances in CRISPR/Cas9 Genome Editing for the Treatment
of Muscular Dystrophies
Sina Fatehi,
1,2
Ryan M. Marks,
1,2
Matthew J. Rok,
1,2
Lucie Perillat,
1,2
Evgueni A. Ivakine,
1,3,{
and Ronald D. Cohn
1,2,4,5,
*
,{
1
Program in Genetics and Genome Biology, Peter Gilgan Centre for Research and Learning, The Hospital for Sick Children Research Institute, Toronto, Ontario,
Canada; Departments of
2
Molecular Genetics and
3
Physiology, University of Toronto, Toronto, Ontario, Canada;
4
Department of Pediatrics, The Hospital for Sick
Children, Toronto, Ontario, Canada; and
5
Institute of Medical Science, University of Toronto, Toronto, Ontario, Canada.
{
Co-senior authors.
Muscular dystrophies (MDs) comprise a diverse group of inherited disorders characterized by progressive muscle loss
and weakness. Given the genetic etiology underlying MDs, researchers have explored the potential of clustered regularly
interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein 9 (Cas9) genome editing as a therapeutic
intervention, resulting in significant advances. Here, we review recent progress on the use of CRISPR/Cas9 genome
editing as a potential therapy for MDs. Significant strides have been made in this realm, made possible through
innovative techniques such as precision genetic editing by modified forms of CRISPR/Cas9. These approaches have
shown varying degrees of success in animal models of MD, including Duchenne MD, congenital muscular dystrophy
type 1A, and myotonic dystrophy type 1. Even so, there are several challenges facing the development of CRISPR/Cas9-
based MD therapies, including the targeting of satellite cells, improved editing efficiency in skeletal and cardiac muscle
tissue, delivery vehicle enhancements, and the host immunogenic response. Although more work is needed to advance
CRISPR/Cas9 genome editing past the preclinical stages, its therapeutic potential for MD is extremely promising and
justifies concentrated efforts to move into clinical trials.
Keywords: muscular dystrophy, Duchenne muscular dystrophy, genome editing, CRISPR/Cas9, adeno-associated
viruses
INTRODUCTION
Muscular dystrophy
MUSCULAR DYSTROPHIES (MDs) are a collection of clini-
cally and genetically heterogeneous disorders character-
ized by progressive muscle weakness and loss.
1,2
The
disorders vary according to the affected muscles, age of
onset, severity, and rate of progression. Currently, most
medical interventions are limited to symptom manage-
ment and delaying disease progression.
1,3
Here, we provide a comprehensive overview of the
current state of genome editing strategies using clustered
regularly interspaced short palindromic repeats (CRISPR)
and CRISPR-associated protein 9 (Cas9) in models of MD.
This work focuses on the utilization of CRISPR/Cas9 as an
in vivo therapeutic strategy to treat Duchenne muscular
dystrophy (DMD), congenital muscular dystrophy type 1A
(MDC1A), and myotonic dystrophy type 1 (DM1). In
addition, we discuss current in vivo delivery options, fo-
cusing on adeno-associated viral (AAV) vectors, recent
advances made, challenges that lie ahead for the field, and
what issues must be resolved for CRISPR/Cas9 genome
editing to become a viable therapy.
DMD is the most prevalent pediatric neuromuscular
disorder due to its X-linked recessive inheritance.
4
It is a
*Correspondence: Dr. Ronald D. Cohn, Program in Genetics and Genome Biology, Peter Gilgan Centre for Research and Learning, The Hospital for Sick Children Research
Institute, 686 Bay Street, Toronto, ON M5G 0A4, Canada. E-mail: ronald.cohn@sickkids.ca
388 jHUMAN GENE THERAPY, VOLUME 34, NUMBERS 9 and 10 DOI: 10.1089/hum.2023.059
ª2023 by Mary Ann Liebert, Inc.
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life-limiting condition caused by mutations in the DMD
gene, which encodes for dystrophin.
5,6
Dystrophin is a
subsarcolemmal protein integral to the dystrophin-
associated protein complex, which prevents contraction-
induced muscle damage.
7
Most commonly, mutations that
cause DMD generate out-of-frame transcripts, eliminating
dystrophin expression.
8,9
MDC1A is an autosomal recessive neuromuscular
disorder characterized by hypotonia, muscle weakness,
and muscle wasting that begins in infancy.
10
MDC1A is
caused by mutations in the LAMA2 gene, which encodes
laminin-2.
11
Laminins are proteins of the extracellular
matrix that are essential components of the basement
membrane.
11,12
DM1 is the most prevalent adult-onset form of MD.
13
Patients may also develop intellectual impairment, respi-
ratory insufficiency, and cardiac conduction abnormali-
ties, in addition to muscle weakness and stiffness.
14,15
DM1 is caused by a microsatellite expansion of cytosine,
thymine, and guanine (CTG) triplet repeats in the 3¢un-
translated region (UTR) of the dystrophia myotonic pro-
tein kinase (DMPK) gene.
13,15
As a result, these mutated
DMPK transcripts generate nuclear foci, which cause ab-
errant splicing across a broad spectrum of pre-mRNAs.
14
Various treatments for MDs are currently approved or
are in clinical testing. Included in this category are anti-
sense oligonucleotides (ASOs), gene therapies, and stop
codon read-through drugs.
1
Although effective in treating
other neuromuscular disorders such as spinal muscular
atrophy (SMA), unfortunately, with regard to MDs, their
relatively poor performance and transient nature limit their
clinical applications.
1
An effective treatment must target
and, ideally, permanently repair the genetic cause of MDs.
Interventions based on CRISPR/Cas9 are extremely
promising because of their unrivalled utility and accuracy
in performing targeted genome editing.
CRISPR/CAS9 GENOME EDITING STRATEGIES
TO TREAT MDS
Exon skipping and reframing
Exon skipping using CRISPR/Cas9 endonucleases can
restore the open reading frame (ORF) of an out-of-frame
gene and lead to the restoration of a functional protein.
Exon skipping has been achieved in DMD patients with the
use of ASOs, which omit the out-of-frame exon(s) from the
final transcript.
16
Eteplirsen, Golodirsen, and Viltolarsen
are examples of such drugs that use ASOs to target out-of-
frame exons in the DMD gene, excluding them from the
final transcript and restoring the reading frame.
17–21
Un-
fortunately, ASOs typically result in low uptake in cardiac
tissue, limiting their clinical relevance regarding the treat-
ment of MDs with a cardiac pathology.
20,21
As a result, CRISPR/Cas9 is currently under investi-
gation as an alternative to mediating exon skipping and
reframing. CRISPR/Cas9-mediated exon-skipping would
be a one-time therapeutic option that results in permanent
correction and restoration of a functional gene product (a
summary of such studies is listed in Table 1). Three main
exon-skipping and reframing approaches (Fig. 1A) have
been tested in preclinical studies and are described next.
Exon removal. This approach aims at restoring the
ORF of a gene by excising the out-of-frame exon(s), re-
sulting in the expression of a truncated protein. This
strategy has been assessed pre-clinically in several DMD
models, including mice, pigs, and dogs.
22–25
Studies in
mice focused on several exons of the Dmd gene, including
exons 23 and 52–53.
23,25
These studies show restoration of
a functional dystrophin protein and improved disease
phenotypes. Studies in pigs and dogs provided proof-of-
principle that CRISPR/Cas9-mediated exon removal is a
viable therapeutic option in larger animal models, paving
the way for its translation to the clinical trials.
22,24
Exon skipping and inclusion through splice site
manipulation. The next strategy designed to mediate
exon skipping relies on manipulation of splice sites using
CRISPR/Cas9 systems. Here, a single-guide RNA
(sgRNA) is used to disrupt splice sites through the error-
prone non-homologous end-joining (NHEJ) repair path-
way, which subsequently allows for the skipping of the
desired exon(s).
26
The NHEJ-mediated introduction of
random deletions and insertions (indels) renders the splice
site no longer identifiable by the spliceosome.
27,28
The disruption results in the exclusion of the exon from
the final transcript, restoring the ORF and expressing a
truncated gene product. This strategy has been tested pre-
clinically in mouse and canine DMD models and resulted
in restoration of a functional dystrophin protein in skeletal
(including the diaphragm) and cardiac tissues.
22,26
In an MDC1A mouse model, Kemaladewi et al. used a
pair of sgRNAs to remove a mutated splice site in the
LAMA2 gene and allowed NHEJ to restore a functional
splice donor site. As a result, full-length laminin-2 was
restored in muscles and the sciatic nerve. Treated mice
exhibited reduced fibrosis, increased muscle fiber size, and
functional improvements comparable to wild-type mice.
29
Indel-mediated reframing. An alternative refram-
ing approach consists of restoring the ORF by introducing
small indels in the coding sequence of an out-of-frame
exon. The Olson group has tested this strategy by using
the engineered KKH variant of SaCas9 in patient-derived
induced pluripotent stem cells (iPSCs) to induce a single
cut that results in the introduction of indels that are able
to reframe exon 51 of the DMD gene.
30
This approach
resulted in a high frequency of indels introducing a two-
nucleotide deletion and, thus, the restoration of the
reading frame.
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Table 1. Summary of in vivo CRISPR studies for the treatment of muscular dystrophies
Reference Model Cas System Strategy Age Vector Route of Delivery Highlights
Exon skipping and reframing
Amoasii et al.
22
Beagle DmdD50 SpCas9 nuclease Exon 51 skipping
through splice site
manipulation and
reframing
4 weeks AAV9 Intramuscular (cranial tibialis)
and systemic (intravenous)
Systemic injection of a high viral dose restored dystrophin expression
between 3% and 92% across all analyzed muscles.
Nelson et al.
23
mdx mouse SaCas9 nuclease Removal of exon 23
using two sgRNAs
P2, 6 weeks AAV8 Intramuscular (tibialis anterior)
and systemic (intraperitoneal)
Intramuscular and systemic injections led to dystrophin expression in
myofibers, along with localization of DAPC proteins and nNOS activity
at the sarcolemma, increasing muscle function.
Amoasii et al.
26
Mouse DmdD50 SpCas9 nuclease Exon 51 skipping
through splice site
manipulation and
reframing
P4, P12 AAV9 Intramuscular (tibialis anterior)
and systemic (intraperitoneal)
Systemic application restored dystrophin protein expression in all
examined muscles. Four weeks after injection, treated mice had
enhanced grasp strength.
Kemaladewi
et al.
29
dy
2J
/dy
2J
mouse SaCas9 nuclease Restoration of splice
donor site
P2, 3 weeks AAV9 Intramuscular (tibialis anterior)
and systemic (intraperitoneal
and temporal vein)
Treated mice exhibited reduced fibrosis, increased muscle fiber size, and
improved function.
Muscle and the sciatic nerve were effectively modified by the systemic
application.
Zhang et al.
30
Mouse DmdD50 SaCas9 (KKH)
nuclease
Exon 51 reframing P4 AAV9 Systemic (intraperitoneal) Systemic delivery of the KKH system restored 30–55% of wildtype levels
of dystrophin in skeletal and cardiac muscles.
Min et al.
31
Mouse DmdD43,
DmdD45 and
DmdD52
SpCas9 nuclease Exon skipping through
splice site
manipulation and
reframing
P12 scAAV9
and ssAAV9
Intramuscular (tibialis anterior) All three mutations were successfully corrected with the single-cut
strategy and resulted in systemic expression of dystrophin between
14% and 23% of wild-type levels.
Hakim et al.
119
GRMD, WCMD, and
LRMD
SaCas9 and
SpCas9 nuclease
Exon removal and exon
skipping through
splice site
manipulation
Newborns,
2–6 weeks
AAV8 Intramuscular (biceps femoris,
cranial tibialis,
semitendinosus, lateral
gastrocnemius, deltoid, and
extensor carpi ulnaris) and
systemic injection (cephalic
vein)
Evaluated intramuscular and intravenous delivery of the Cas9 editing
system using AAV8s in three canine models of DMD. Demonstrated
that AAV-mediated expression of CRISPR reagents induced a humoral
and B and T cell-based immune response to Cas9, which reduced the
extent and duration of dystrophin restoration.
Min et al.
127
Mouse DmdD44 SpCas9 nuclease Exon 43 and 45 skipping
through splice site
manipulation and
reframing
P12 AAV9 Intramuscular (tibialis anterior)
and systemic (intraperitoneal)
Cas9 editing is dependent on the quantity of sgRNA present; therefore,
sgRNA is the limiting factor for in vivo editing efficacy. Systemic
administration of 1:10 Cas9:sgRNA led to 90–95% of wild-type
dystrophin expression in cardiomyocytes and myofibers.
Hakim et al.
128
mdx mouse SaCas9 nuclease Removal of exon 23
using two sgRNAs
6 weeks AAV9 Intramuscular (tibialis anterior)
and systemic injection (tail
vein)
Evaluated long-term AAV-mediated CRISPR editing in mdx mice and
observed significant dystrophin restoration in skeletal and cardiac
muscles. Demonstrated disproportional sgRNA vector depletion.
Increasing the sgRNA vector ameliorated the issue, enhancing muscle
histology and hemodynamics.
Young et al.
129
hDMD/mdx and
hDMDD52/mdxD2
SpCas9 nuclease Removal of exon 45–55
using two sgRNAs
12–18.5 weeks Plasmids Muscle electro-poration of flexor
digitorum brevis
First investigation using the CRISPR system to target hDMD/mdx model.
(continued)
390 j
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Table 1. (Continued)
Reference Model Cas System Strategy Age Vector Route of Delivery Highlights
Duplication removal
Maino et al.
38
Exons 18–30 duplication SaCas9 nuclease Single sgRNA
duplication removal
P1–P2 AAV9 Systemic (temporal vein) Treated mice demonstrated significant dystrophin protein restoration in
various muscle groups, notably the heart, averaging 17% of wildtype
levels. The dystrophic phenotype was significantly improvement based
on histological and functional assays.
Transcriptional modulation of disease modifiers in MDs
Kemaladewi
et al.
56
dy
2J
/dy
2J
mouse SaCas9 nuclease Upregulation of Lama1 P2, 3 weeks AAV9 Intramuscular (tibialis anterior)
and systemic injection
(temporal vein)
Treatment of aged, symptomatic mice improved and effectively reversed
the progression of the disease.
Pinto et al.
71
HSA
LR
mouse carrying a
human skeletal actin
transgene containing
250 CTG repeats
dSpCas9 nuclease Transcriptional
inhibition of
expanded
microsatellite repeats
P2 AAV6
and AAV9
Systemic (temporal vein) First published study inhibiting transcription in a DM1 murine model using
dCas9. 5–15% of fibers exhibited total absence of CUG RNA foci.
Base-editing in MDs
Ryu et al.
84
Dmd exon 20
A-T to G-C point
mutation
ABE7.10-nSpCas9 Remove premature
termination codon
7 weeks tsAAV9 Intramuscular (tibialis anterior) Restoration of dystrophin expression via modifying a premature
termination codon. Restored expression of dystrophin in *15%
myofibers.
Chemello et al.
85
Mouse DmdD51 ABEmax-nSpCas9-NG Exon-skipping via splice-
site disruption
P12 Split-intein
AAV9
Intramuscular (tibialis anterior) Exon-skipping via disruption of exon splicing motifs. Restored expression
of dystrophin in *96.5% myofibers.
Arbab et al.
86
Smn2D7C6T ABE8e-SpMAC
+nusinersen
6TAto6CG via
adenine base editing
P0 Split-intein
AAV9
Intracerebroventricular Demonstrated the utility of combinatorial approaches in genetic medicine
(ASO+base editing) for synergistic therapeutic outcomes. Extended the
therapeutic window for base editing via early pharmacologic
intervention.
CRISPR/Cas9-mediated exogenous DNA knock-in to restore wild-type proteins
Bengtsson
et al.
25
mdx
4cv
mice with non-
sense mutation in
exon 53
SaCas9 and
SpCas9 nucleases
Deletion of exons 52–53
and exogenous DNA
knock-in (HDR)
10–12 weeks AAV6 Intramuscular (tibialis anterior)
and systemic (retro-orbital)
Removed exon 53 while delivering an HDR template to correct the
mutation. Retro-orbital delivery of AAV6s carrying CRISPR/Cas9
nucleases yielded *34% dystrophin expression in the heart as well as
widespread expression in all analyzed muscle tissues. HDR correction
efficiency was measured at 0.18%.
Mata Lo
´pez
et al.
92
GRMD with a mutation
in intron 6 splice
acceptor site
SpCas9 nuclease Exogenous DNA knock-
in (HDR)
3 months–8
years
Plasmid Intramuscular (cranial tibialis,
long digital extensor and
peroneus longus)
Intramuscular injections of the CRISPR/Cas9 treatments resulted in
inclusion of exon 7 in the transcript, dystrophin restoration between at
2% and 16%.
Lee et al.
93
mdx mouse SpCas9 RNP Exogenous DNA knock-
in (HDR)
4 weeks Gold
nano-particles
Intramuscular (tibialis anterior
and gastrocnemius)
Simultaneous treatment with CRISPR-Gold and cardiotoxin led to
widespread dystrophin restoration with HDR efficiency of 5.4%.
Pickar-Olive
et al.
96
hDMDD52/mdx SaCas9 nuclease Exogenous DNA knock-
in (HITI)
P2 neonates AAV9 Intramuscular (tibialis anterior)
and systemic (temporal vein)
First study to devise a DNA knock-in strategy around NHEJ. Systemic
delivery of components using a dual AAV9 system was able to achieve
knock-in rates of 4.67–7.47%. The ‘‘superexon’’ strategy could
potentially correct a large of known DMD mutations.
AAV, adeno-associated virus; ASO, antisense oligonucleotide; Cas9, CRISPR-associated protein 9; CRISPR, clustered regularly interspaced short palindromic repeats; CTG, cytosine, thymine, and guanine; DAPC, dystrophin-
associated protein complex; dCas9, dead Cas9; DM1, dystrophy type 1; DMD, Duchenne muscular dystrophy; GRMD, golden-retriever model of DMD; HDR, homology-directed repair; HITI, homology-independent targeted
integration; LRMD, Labrador retriever muscular dystrophy; MD, muscular dystrophy; NHEJ, non-homologous end-joining; RNP, ribonucleoprotein; sgRNA, single-guide RNA; tsAAV, trans-splicing adeno-associated virus; WCMD,
Welsh corgi muscular dystrophy.
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The group subsequently tested this approach in vivo in
mice with a deletion of Dmd exon 50. Results demon-
strated that this single-cut strategy restored dystrophin
expression to 30–50% in skeletal muscle and 50% in the
heart, along with improved muscle integrity and function.
The authors note that using SpCas9 tends to favor NHEJ-
mediated insertion of one base pair, which is not the case
with SaCas9. To address this challenge, the authors
screened for guide RNAs that can induce double-strand
breaks (DSBs) in regions of microhomology, which tend
to produce predictable deletions.
Min et al. also tested single-cut genome editing strate-
gies in mouse and cell models of DMD.
31
They compared
both reframing and exon skipping for three mutations
Figure 1. Summary of in vivo validated CRISPR/Cas9 genome editing strategies for the treatment of MDs. On the left are gene stretches containing a mutated
exon, highlighted in red, as well as respective CRISPR/Cas9 strategies, leading to the desired outcomes on the right.(A) Restoring the ORF through CRISPR/
Cas9-mediated exon deletion, exon skipping, and exon reframing. (B) Removal of a duplicated region using single and dual sgRNA approaches. (C) Modulating
the expression of a disease modifier using CRISPR/dCas9 fused to a transcriptional activator or repressor. (D) Correction of a splice site mutation (A–G) using
an adenine BE. (E) HDR and NHEJ mediated exogenous DNA knock-in of a deleted exon. BE, base editor; Cas9, CRISPR-associated protein 9; CRISPR,
clustered regularly interspaced short palindromic repeats; dCas9, dead Cas9; HDR, homology-directed repair; MD, muscular dystrophy; NHEJ, non-homologous
end-joining; ORF, open reading frame; sgRNA, single-guide RNA.
392 FATEHI ET AL.
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using SaCas9. From their results, they concluded that +1
reframing has comparable editing efficiencies to -1 re-
framing depending on a chosen guide RNA. They also
note that the indel type observed in iPSC-derived cardio-
myocytes can be used to predict the levels of dystrophin
restoration in vivo.
Advantages, limitations, and future directions of
CRISPR/Cas9-mediated exon skipping and re-
framing. CRISPR/Cas9-mediated exon skipping and
reframing has several advantages compared with ASOs,
including the ability to potentially achieve a permanent
correction using a single treatment, which significantly
improves the cost-benefit profile of this approach, espe-
cially as the cost of clinical-grade AAV production con-
tinues to decrease.
32
Exon skipping and reframing have
been tested in vitro and in vivo, including in large animal
models, and have led to significant and widespread res-
toration of dystrophin. The major limitation of these
strategies is that exon skipping and reframing typically
lead to the production of a truncated gene product. De-
pending on which exons are skipped or lost, this may lead
to varying clinical improvements.
33
To address this, focus
should be placed on studying the impact of such correction
strategies on patients as we approach clinical trials.
Duplication removal
Large genomic duplications typically encompass one or
more exons and result in reading frame disruption or im-
pairment of protein function through the addition of these
exons to the transcript. In DMD, these duplications rep-
resent 10–15% of patient mutations and occur at low but
appreciable frequencies for several other MDs, such as
MDC1A.
34–37
Therapeutics specific for treating MD du-
plication mutations have been mostly neglected, leading to
a significant unmet need for a sizeable patient population.
Presently, applications of CRISPR/Cas9 toward treating
large duplications have only been published for DMD,
though the principles behind these studies should be
broadly applicable to other MDs.
Duplication removal using single- and dual-sgRNA
approaches. Large duplications can be corrected in
one of two ways using CRISPR/Cas9 (Fig. 1B). The first
employs two sgRNAs that flank the duplicated region and,
through coordinated cleavage, excises the intervening
duplication, with subsequent ligation restoring the wild-
type sequence. The second relies on a solitary sgRNA
targeting within the duplicated sequence. As a result, the
original and duplicated sequences are cut in the same lo-
cation with the intervening sequence being excised, also
restoring the wild-type sequence.
There are several key advantages to the latter approach
that make it the preferred option for correcting most large
duplications. Using one sgRNA rather than two notably
reduces the complexity of delivery. Minimizing the es-
sential CRISPR components is critical, as AAVs will
likely remain the delivery vehicle of choice for clinical
trials for the foreseeable future. The single-sgRNA ap-
proach enables packaging of all components into one
AAV.
38
Using the same sgRNA for both target sites may
improve coordinated cutting at both sites by reducing the
potential for asynchronous cleavage and thus enhance
excision of the duplication. In addition, the entire dupli-
cated region is made available for guide selection, which
enables the assessment of numerous sgRNAs to identify
the most efficacious candidates. It should be noted that to
minimize potential disruption to the coding sequence,
sgRNA design should be limited to introns.
In DMD patients, large duplications typically manifest in
tandem, which enables the use of the single-sgRNA ap-
proach. It is worth noting that non-contiguous DMD dupli-
cations have been reported, whereby a dual sgRNA strategy
may be the only option.
39,40
Several groups have demon-
strated the successful restoration of full-length dystrophin in
DMD patient-derived myoblasts harboring large duplica-
tions using the solitary sgRNA strategy. Wojtal et al. were
the first in 2016, correcting a duplication of exons 18–30.
41
Studies soon followed, correcting DMD patient dupli-
cations of exon 2, exons 55–59, exons 18–25, and exons 3–
16.
42–45
We were unable to identify any published study
utilizing the dual sgRNA strategy for correcting a dupli-
cation mutation, likely owing to its notable disadvantages
and the relatively early reporting of the successful use of a
solitary sgRNA. Unfortunately, in vivo application of
CRISPR/Cas9 for correcting DMD duplications is limited
to a single 2021 study by Maino et al.
There, a novel DMD mouse model harboring a 130 kb
exon 18–30 duplication was generated and subsequently
treated with the single-sgRNA approach using SaCas9 and
delivered systemically by AAV9s. Significant full-length
dystrophin restoration, averaging *17%, was observed in
muscle tissues, most notably in the heart, resulting in sig-
nificant improvements in muscle histology and function.
38
Advantages, limitations, and future directions of
CRISPR/Cas9-mediated duplication removal. In vivo
studies on large duplications are extremely limited owing
to the immense difficulty in generating appropriate animal
models relative to large deletions and small insertions or
duplications. In addition to the exon 18–30 duplication
mouse, only one other model has been described, which
possesses an exon 2 duplication, the most common du-
plication observed in DMD patients.
39,46
However, this
model was obtained by inserting exon 2 and only small
portions of its flanking intronic sequences into a duplica-
tion hotspot in intron 2.
46
As a result, the full genomic architecture of a typical
large tandem duplication is lost, limiting investigations
of duplication removal strategies. There does remain
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immense utility for this model, as demonstrated by the
successful evaluation of an AAV-packaged ASO, which is
currently under investigation in a clinical trial.
47–49
Most
recently, a family of Labrador retrievers possessing a
tandem 400 kb duplication of exons 2–7 in the DMD gene
has been described and may prove to be an important large
animal model for pre-clinical research into duplication-
specific CRISPR/Cas9 therapies.
50
Although ASOs demonstrate some potential use for
single exon duplications, care must be taken as different
exons respond variably to ASOs, with non-productive
skipping of both exons having been observed.
40,47,51
Multi-exonic duplications magnify the difficulty propor-
tional to their size and remain extremely problematic to
treat with ASO.
51
CRISPR/Cas9 provides a solution for
treating duplications, particularly tandem ones, using
single- and dual-sgRNA approaches. Maino et al. have
importantly demonstrated that generating mouse models
with large multi-exonic duplications is feasible.
38
Looking ahead, focus should be shifted toward devel-
oping novel duplication animal models of various MDs
that faithfully reflect patient genomic architecture, in-
cluding humanized mouse models. These will be essential
for enabling translational CRISPR/Cas9 preclinical re-
search. Although studies on the topic remain sparse, there
is a growing appreciation for large duplications in MD and
how CRISPR/Cas9 can be applied to correct them.
Transcriptional modulation of disease
modifiers in MDs
CRISPR/Cas9 technology can be harnessed to modulate
the expression of target genes of interest (Fig. 1C). In this
approach, it is typical to target an inactivated or ‘‘dead’’
Cas9 (dCas9) protein, fused to a transcriptional activator or
repressor, to the promoter region of a target gene.
52
CRISPR activation (CRISPRa) enables overexpression of
target genes via recruitment of transcriptional machinery or
relaxation of surrounding chromatin.
53
CRISPR interference (CRISPRi) is a similar technique
that enables the suppression of target gene expression by
interfering with transcriptional machinery to transiently
inhibit gene expression.
54
The technique works by tar-
geting a dCas9 fused to a transcriptional repressor, to the
promoter region of the gene of interest. Alternatively,
dCas9 can be fused to a variety of epigenetic enzymes that
perform biochemical modifications to condense local
chromatin structure and/or methylate promoters, masking
the gene of interest from transcriptional machinery. For a
more detailed description of CRISPR/Cas9-mediated
transcriptional regulation systems, please refer to the fol-
lowing review.
55
CRISPR/Cas9-mediated transcriptional regulation
of MD disease modifiers. Disease modifier genes are
those, separate from the mutated gene, that can affect the
severity and progression of a disorder. Restoring full-
length dystrophin via gene therapy remains challenging
due to its large size.
1
Expression of alternative homologs
capable of performing the functions of dystrophin would
be an ideal mutation-independent approach, applicable to
all patients.
56,57
Utrophin is a cytoskeletal protein, highly
similar in structure and function to dystrophin, encoded by
the autosomal UTRN gene.
58–61
Utrophin is primarily expressed in fetal skeletal muscle
and is largely replaced by dystrophin in adult tissues.
58
Interestingly, patients with DMD have endogenously in-
creased levels of utrophin distributed in damaged muscle,
and several early studies demonstrated that utrophin
overexpression could rescue the DMD phenotype in
mice.
62,63
In fact, double knockout mice of Dmd and Utrn
possess a far more severe phenotype than Dmd null
mice.
64,65
Although several small molecule-based ap-
proaches are in development to promote utrophin ex-
pression and/or decrease the rate of UTRN transcript
downregulation,
59,66,67
CRISPRa has been demonstrated to be effective as a
highly specific and efficacious method for upregulation of
utrophin. Wojtal et al. demonstrated that a dCas9-VP160
fusion targeting either the UTRN A or Bpromoters could
effectively upregulate utrophin protein expression in
patient-derived myoblasts.
41
Although these results were
recently validated in DMD patient-derived stem cells,
57
this approach requires assessment of efficacy, longevity of
utrophin expression, and characterization of its effects on
the dystrophic phenotype in vivo.
A similar CRISPRa approach has been used to treat
MDC1A via disease modifier upregulation. There are over
350 known pathogenic nonsense, missense, splice site, and
deletion mutations in LAMA2; an efficacious disease
modifier would provide a therapeutic alternative to gene-
editing that is applicable to all MDC1A patients.
68
LAMA1, which encodes the structurally similar laminin-a1
protein, was demonstrated via overexpression studies to be
a suitable candidate to substitute the function of laminin-
a2 in vivo.
Kemaladewi et al. were able to upregulate Lama1 in
skeletal muscle via systemic delivery of a dSaCas9-VP64
protein in mice harboring a splice site mutation in the
Lama2 gene.
56
This strategy led to significant expression
of the laminin-a1 protein and was correlated with im-
provements in the histological hallmarks of MDC1A, in-
cluding significantly reduced fibrosis. Importantly,
systemic treatment of MDC1A mice led to a remarkable
rescue of the advanced hind-limb paralysis phenotype,
with significant improvements in ambulation as well as
increases in specific tetanic force and conduction velocity
in the sciatic nerve.
It is predicted that this approach may also provide new
opportunities for treatment of DMD, as overexpression of
Lama1 has been shown to stabilize the sarcolemma in
394 FATEHI ET AL.
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dystrophic myofibers from mice with dystrophinopathy,
further expanding the utility of this approach.
69,70
Ulti-
mately, these results demonstrate the utility of disease
modifiers as legitimate therapeutic strategies for the
treatment of MDs.
The modular nature of the CRISPR/dCas9 system
means it can be used not only to upregulate but also to
downregulate the expression of a target gene. Pinto et al.
treated the transgenic HSA
LR
mouse, which carries a
fragment of the human skeletal actin (HSA) gene with 250
CTG repeats in the 3¢UTR, with dCas9 targeting the CTG
repeats and preventing their inclusion in HSA tran-
scripts.
71
This resulted in a significant decrease in repeat
transcription and an improvement in phenotype. Appli-
cations involving a combinatorial approach with concur-
rent upregulation of protective disease-modifier genes and
downregulation of detrimental genes could represent a
new mutation-independent approach for ameliorating
disease phenotype.
Advantages, limitations, and future directions of
CRISPR/Cas9-mediated transcriptional regulation of
disease modifiers. Modulation of gene expression by
CRISPR has many advantages over traditional nuclease-
based approaches. First, as CRISPRa/CRISPRi is based on
fusions of effector domains linked to a catalytically in-
activated Cas9, there is negligible risk for the introduction
of double-stranded DNA breaks at the target site and the
induction of severe chromosomal damage.
72,73
Although it is still possible for a dCas9-fusion to lo-
calize effector modules to an off-target location in the
genome, it is unlikely that this region is proximal to the
promoter or enhancer motifs, further reducing the chance
of modulating the transcriptional output of an off-target
gene.
74
Transcriptomic studies from the dCas9-VP160-
mediated Lama1 upregulation study largely suggest this
concern is negligible.
56
Second, modulation of disease
modifiers via CRISPRa/CRISPRi offers a mutation-
independent approach to treating genetic diseases.
Combining CRISPR transcriptional modulation with
traditional CRISPR-mediated gene editing strategies can
generate novel therapeutic strategies to treat MDs. These
would involve using CRISPRa to increase the expression of
a CRISPR/Cas9-corrected gene. We believe the primary
advantage of this combinatorial strategy may be especially
relevant to diseases where a small number of corrected
nuclei can rescue a phenotype by supplying sufficient gene
product for the entire tissue. MDs can also benefitfrom such
a strategy since in many neuromuscular disorders, a single
edited nucleus can complement the entire myofiber syn-
cytium with a functional gene product.
75
Base-editing in MDs
The introduction of base-editing was the first iteration
of true gene-editors capable of producing targeted modi-
fications to specified regions of DNA (Fig. 1D).
76
Base-
editing directly generates precise point mutations in
genomic DNA without double-stranded breaks, a DNA
donor template, or cellular homology-directed repair
(HDR).
76–78
These editors comprised a fusion between a
catalytically impaired Cas9 (D10A) nickase and a base-
modifying enzyme that operates on transient single-
stranded DNA intermediates.
76,79
Two classes of DNA base editors have been described
to date: cytosine base editors (CBEs), which convert a C-G
base pair into a T-A base pair, and adenine base editors
(ABEs), which convert an A-T base pair into a G-C base
pair.
76,80
Collectively, CBEs and ABEs can mediate
all four possible transition mutations (C-to-T, A-to-G,
T-to-C, and G-to-A). More information can be found on
the development of base editors in this review by Rees
and Liu.
81
Use of base-editors in the therapeutic context for
MDs. Therapeutic applications of base editing for
neuromuscular disease are a promising strategy due to its
highly efficient nature and precise editing of pathogenic
point mutations in post-mitotic cells, including neurons
and myofibers.
82
Specifically, base editors have the po-
tential to be used as therapeutic interventions in *16% of
DMD patients who harbor a pathogenic single nucleotide
polymorphism that could be corrected by this technology
alone.
83
Base editors have been utilized as such in two
publications, which highlight their potential for correcting
nonsense mutations and manipulating splice sites in exon-
skipping strategies in vivo.
Ryu et al. used a trans-splicing adeno-associated virus
vector system to deliver an ABE7.10 base editor that was
programmed to target a point mutation in Dmd exon 20, a
causative mutation that introduces a premature termina-
tion codon and subsequently produces a DMD pheno-
type.
84
Intramuscular delivery of the ABE7.10 via AAV9s
to the tibialis anterior (TA) was able to successfully per-
form single-nucleotide substitution via A-T to G-C de-
amination to restructure the premature termination codon
at a frequency of 3.3%, 8 weeks post-injection.
Notably, this genomic correction was accompanied by a
17% restoration of dystrophin-positive myofibers com-
pared to control. Base editors can also be used as a mod-
ified approach for exon-skipping in DMD, as described
earlier. Chemello et al. utilized a dual AAV9 split-intein
trans-splicing ABEmax-SpCas9-NG system to accomplish
exon-skipping via A-T to G-C deamination of the splice
donor site to interrupt it and exclude exon 50 from the
mature Dmd transcript in a Dmd exon 51 deletion mouse.
85
Intramuscular injection into the TA achieved an on-
target genomic editing efficiency of 35%, and Sanger se-
quencing of transcripts illustrated scarless splicing of Dmd
exon 49 to exon 52 as expected. Immunoblotting for full-
length dystrophin demonstrated 54% protein restoration
CRISPR/CAS9 TREATMENT OF MUSCULAR DYSTROPHIES 395
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compared with wild-type control. Histological findings
suggested a substantial reduction in fibrosis and 96.5%
restoration of dystrophin-positive fibers in treated animals.
However, the authors note that the titer of intramuscular
AAV9s delivered in this study would extrapolate to a
systemic intravenous dosage corresponding to 1.5E16 vi-
ral genomes per milliliter, significantly higher than an
acceptable threshold for therapeutic usage.
In a recent study, the Liu lab used an ABE to treat an
SMA mouse model (D7SMA).
86
Conveniently, SMN1 has
a paralogous gene in humans known as SMN2, which
mainly differs from SMN1 by a C-G-to-T-A transition in
exon 7, resulting in the skipping of exon 7 in most SMN2
transcripts and the production of only low levels of SMN.
Following intracerebroventricular injection of a split-
intein ABE-Cas9 packaged in an AAV9 vector in neonatal
mice, the authors could detect *40% A-to-G editing on
average in bulk cortical tissue. The treatment also in-
creased the lifespan of the D7SMA mice by *33%, al-
though the authors note that the short lifespan of these
animals makes phenotypic analysis challenging.
Therapeutic advantages, limitations, and future
directions of base-editing in MDs. Although the
technological framework has been rapidly developed to
date, there exist several key challenges to the translation of
base-editing into the clinic, some of which are currently
being resolved. First, the original base-editing platforms
are too large for packaging into traditional AAV vectors
in clinical use today.
87
Currently, most preclinical
base-editing work is being performed using a dual-vector
system, whereby the base editor is assembled post-
translationally in a co-transduced cell via a split-intein
mechanism.
85,88
The nature of dual-vector systems necessitates in-
creased titers of viral vehicles, which possess immuno-
logical and toxicological concerns. In addition, dual
vector systems are more expensive to produce. None-
theless, the Liu lab has recently developed an optimized
suite of ABEs that can be constructed into a single, all-in-
one vector with the added benefit of increased efficiency
of transgene expression, suggesting potentially lower
titer requirements.
89
This was achieved by fusing a single evolved TadA
deaminase domain to smaller nickase variants of Cas9
(SaCas9, SaKKH Cas9, SauriCas9, and NmeCas9), which
has the additional benefit of creating a toolbox of base
editor enzymes with alternative or expanded protospacer
adjacent motif or PAM, theoretically permitting base-
editing at *80% of sites within the genome by the com-
bination of these four smaller nickases. In addition, the
group has utilized directed evolution to generate highly
active cytosine deaminase enzymes from the previously
evolved and smaller adenosine deaminase, as well as hy-
brid deaminase enzymes capable of both adenosine and
cytosine deamination. These novel base editor variants
should lead to the development of CBEs that are small
enough for single AAV packaging in the near future.
CRISPR/Cas9-mediated exogenous DNA
knock-in to restore wild-type proteins
To permanently correct MDs, restoration of the wild-
type protein at physiologically relevant levels is required.
Multi-exonic deletions constitute the majority of DMD
patient mutations (*60%). For precise CRISPR/Cas9
correction of deletions, two DNA knock-in strategies have
been pursued: HDR and NHEJ (Fig. 1E).
HDR-mediated exogenous DNA knock-in. Al-
though HDR is not an efficient pathway in muscle cells
due to their post-mitotic nature,
90,91
several investigations
have attempted to correct a variety of mutations using this
strategy in vivo. In a golden-retriever model of DMD,
Mata Lo
´pez et al. demonstrated correction of a splice ac-
ceptor mutation by delivering plasmids carrying the Cas9
endonuclease and an HDR donor template to the tibio-
tarsal flexor muscle, restoring 2–16% of wild-type dys-
trophin levels.
92
In another study, local delivery of AAV9s carrying
CRISPR/Cas9 and an HDR template to the TA success-
fully replaced exon 53 with an efficiency of 0.18%.
25
Fi-
nally, Lee et al. obtained an HDR frequency of 0.8% in
mdx mice via local delivery using gold nanoparticles
(CRISPR-Gold).
93
HDR efficiency was raised to 5.4%
when mdx mice were simultaneously treated with cardio-
toxin, which activates the proliferation of muscle stem
cells.
93,94
Although several potential HDR-based strate-
gies have been developed to repair mutations in other
forms of MDs, no in vivo studies have been conducted
to date.
95
The inability of HDR-based techniques to restore
clinically significant quantities of protein in muscle cells
prevents their use as a treatment for MD. These findings
demonstrate the limitations of HDR and the need for al-
ternative strategies to improve its effectiveness in muscle.
NHEJ-mediated exogenous DNA knock-in. NHEJ
is the predominant DNA repair pathway in muscle cells
that corrects DSBs. Unlike HDR, NHEJ does not require a
homologous repair template. Homology-independent tar-
geted integration (HITI) relies on CRISPR/Cas9 and
NHEJ to achieve precise DNA knock-ins. Pickar-Oliver
et al. used HITI to knock-in exon 52 of DMD in a hu-
manized mouse model (hDMDD52/mdx) both locally and
systemically.
96
Localized delivery to the TA using AAV9s
resulted in knock-in rates of 0.24–0.67%. In addition to
reintroducing a single exon, a novel strategy using a
‘‘superexon’’ donor was investigated.
This knock-in template includes DMD exons 52–79 and
a synthetic poly-A signal, which has the potential to
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correct many DMD mutations located downstream of exon
51. Delivery of the superexon to TAs resulted in knock-in
rates of 0.17%. This treatment was also administered
systemically and resulted in knock-in efficiencies of 0.15–
3.05% across multiple tissues, including both skeletal and
cardiac muscles. Although generally more efficient at in-
troducing exogenous DNA sequences to the genome of
muscle cells, NHEJ-based knock-in strategies are still too
inefficient to warrant translation to clinical trials. Further
investigation is required to improve the efficiency of this
approach in skeletal and cardiac tissue.
Advantages, limitations, and future directions of
CRISPR/Cas9-mediated exogenous DNA knock-
Ins. Low editing efficiencies in muscle are a major
hurdle slowing the development of DNA knock-in strat-
egies to treat MDs. Effective integration of exogenous
DNA would enable a range of new CRISPR applications,
including those unrelated to MDs. Due to the post-mitotic
nature of myotubes and satellite cells, alternative strate-
gies to HDR must be developed to allow for efficient
editing in these cells.
97,98
HITI, homology-mediated end
joining (HMEJ), and prime editing are three emerging
techniques that could address this challenge.
96,99,100
It is proposed that HMEJ occurs via single-strand an-
nealing using a modified DNA donor template that con-
tains sgRNA target sites for its excision from a delivery
vector. AAV9 delivery into the mouse cortex resulted in a
transduced neuron population with a knock-in rate of 50%.
An alternative in vitro study on human-derived IPSCs and
myoblasts demonstrated knock-in rates of 6–30% when
inserting the full DMD transcript.
101
HMEJ is a promising
strategy for the correction of a multitude of MD mutations,
although further investigation is required regarding its
efficacy in vivo.
Cas9 nickase and reverse transcriptase proteins are
fused to create prime editors.
100
Utilizing a 3¢-extended
pegRNA encoding the required template, comparable
in vitro knock-in rates to HDR have been achieved. Che-
mello et al. rescued a deletion of DMD exon 51 by de-
livering a prime editor to patient-derived IPSCs.
85
This
strategy aimed at restoring the expression of the gene by
reframing exon 52 through a +2 insertion and achieved a
correction rate of 20.2%. Prime editing, however, is highly
inefficient at introducing larger sequences of exogenous
DNA, such as those the size of exons.
100,102
Novel strategies such as twin-PE allow for the intro-
duction of large sequences into the genome using prime
editors and integrases together.
102
This creates the possi-
bility of designing systems that can integrate entire gene
transcripts, including engineered promoter regions to ex-
press proteins of interest at physiologically relevant levels.
For in vivo MD applications, however, downsizing of the
technology and validation in muscle cells are still required.
It, nevertheless, presents a promising avenue to pursue.
IN VIVO CRISPR/CAS9 DELIVERY SYSTEMS
Progress and current challenges with
adeno-associated viral vectors
Adeno-associated vectors have emerged as a promising
tool for gene therapy in MDs.
103,104
One of the most
promising AAV serotypes for MD gene therapy is AAV9,
which has shown remarkable efficacy in preclinical stud-
ies.
105–108
AAV9 has a broad tropism for various cell
types, including muscle cells, and can efficiently transduce
both skeletal and cardiac tissues.
109
However, AAVs also
have limitations and disadvantages that must be consid-
ered. One of the main disadvantages is their limited cargo
capacity.
109
In addition, AAVs can elicit an immune response,
particularly on repeated administration or due to pre-
existing immunity in some patients.
110–113
Further, the
production and purification of AAV vectors is currently
complex and expensive, leading to high manufacturing
and treatment costs for patients.
32
Despite these chal-
lenges, the use of AAVs for gene therapy is a promising
approach, and ongoing research is focused on overcoming
these limitations and further advancing the clinical ap-
plication of AAV-mediated gene therapies for MDs.
In 2020, the Grimm group described a novel AAV
capsid variant termed AAVMYO, which consistently
outperformed AAV9s in skeletal and cardiac tissue when
analyzing the level of transcribed cargo.
114
Another recent
advance is the generation of a novel muscle-specific se-
rotype termed MyoAAV1.
115
With a novel engineered
capsid, MyoAAV1 promises higher specificity for muscle
tissue and reduced tropism for the liver. Increasing the
efficiency of cargo delivery to skeletal and cardiac cells
while reducing the potential for side effects is associated
with off-target transduction. Further research is needed to
fully evaluate the safety and efficacy of AAVMYO and
MyoAAV1 for the treatment of MDs.
DISCUSSION
Long-term considerations for the use
of in vivo CRISPR gene editing
It is essential to understand the long-term consequences
and effects of utilizing CRISPR-mediated editing as a
therapeutic strategy. The stability of genomic manipula-
tion in muscle cells is still poorly understood within the
context of MD patients, with a key concern being the rate
of muscle turnover. It is our opinion that without the
correction of satellite cell populations, it is unlikely that
the life-long benefits of a single, virally mediated treat-
ment will be realized. This issue is especially pertinent for
MDs, where the optimal therapeutic window is early
childhood, before the progression of more severe symp-
toms.
1,86
Rapid muscle growth at this age may quickly
render CRISPR therapies that target non-satellite cells
ineffective through the loss or dilution of corrected nuclei.
CRISPR/CAS9 TREATMENT OF MUSCULAR DYSTROPHIES 397
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Although AAVs are considered to be a relatively effi-
cient vector for use in humans, adverse effects have been
documented, which may limit their clinical utility. Im-
munogenicity against the virus and the Cas9 machinery, as
well as integration of the AAV into the genome, further
exacerbate immunological and genotoxic con-
cerns.
113,116,117
In a study by the Gersbach group, a hu-
moral and cellular immune response was observed in adult
mdx mice when treated by AAV8/9s carrying an SaCas9,
which was mostly avoided when the same treatment was
given to P2 neonates.
117
AAV-based delivery is the current gold standard for
FDA-approved systemic treatments, and the dose-
dependent adverse effects of AAV, such as temporary
thrombocytopenia and transaminitis, can be effectively
managed.
112,118
The Duan group evaluated intramuscular
and intravenous Cas9 and sgRNA delivery in three
distinct canine models of DMD to measure the effect of
pre-existing Cas9 immunity.
119
They discovered that
AAV-mediated expression of CRISPR reagents not only
promoted editing of the DMD gene but also induced a
B and T cell-based immune response to Cas9, which re-
duced the extent and duration of dystrophin restoration.
Expression of a novel protein that was absent during the
removal of self-reactive lymphocytes can stimulate im-
munological responses.
113
Anti-dystrophin antibodies and
immunological rejections have been observed in DMD
animals who have received remedial treatments.
120
Sev-
eral studies have also demonstrated significantly higher
rates of AAV integration into the genome than previously
thought.
113,116,117
AAV genomes can spontaneously inte-
grate into the AAVS1 safe harbor site on chromosome 19
and were previously thought to rarely integrate into DSBs
created in the genome by Cas9.
121
Across two studies, the Gersbach group reports be-
tween *1and*5% of editing outcomes, resulting in
AAV genome integration in both skeletal and cardiac
tissue when Cas9 nucleases were delivered systemically
to mice.
96,117
AAV integration events can pose serious
immunogenic concerns when paired with pre-existing
Cas9 immunity. Hence, along with pharmacological op-
tions such as immunosuppressants to counter immune
Figure 2. Current challenges with in vivo CRISPR/Cas9 genome editing for the treatment of MDs. An overview of the technical difficulties associated with the
use of AAVs for MD therapies, delivery of CRISPR/Cas9 components to satellite cells, immune responses to targeted or edited cells, and CRISPR/Cas9 editing
efficiencies in muscle. AAVs, adeno-associated viruses.
398 FATEHI ET AL.
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reactions, extensive safety monitoring will be required
during clinical trials to reduce immune response in
patients.
Conclusions and future directions for the MD
gene editing field
Current innovations in the application of CRISPR/Cas9
have created various new treatment prospects for MDs.
There is tangible excitement in this field, but there con-
tinues to be a lot of necessary work ahead as we begin to
consider clinical trials for these therapeutic avenues. Here,
we lay out four areas that require the most focus: efficient
targeting of satellite cells, improved editing efficiency in
muscle tissue, vector enhancements, and immunogenic
response (Fig. 2).
An ideal MD therapy would target satellite cells, and so
it is imperative to determine if currently available serotypes
can target these cells efficiently. If not, further work needs
to be done to design or evolve a novel serotype that can
efficiently deliver gene editing components to these targets.
Other systems, such as virus-like particles or nanoparticles,
are also potential candidates for this role but require further
in vivo testing regarding efficacy and safety.
CRISPR/Cas9 editing efficiencies are lower than desired
when targeting muscle tissue. Although less than ideal rates
of correction in muscle are partly due to the challenges
associated with delivery of editing machinery to cells, there
is a need to further develop more efficient systems and
strategies. Tools such as base and prime editors are prom-
ising but require further development and in vivo validation.
Knock-in strategies allow for restoration of the wild-type
proteins; however, current approaches such as HDR are too
inefficient. Development of systems reliant on HMEJ and
integrases could open new opportunities to develop MD
therapies based on DNA knock-ins.
AAV9s will likely be used in the first generation of
CRISPR therapeutics for MDs, given their track record in
in vivo investigations and clinical trials. AAV9s may be
the best option available, but future therapies will certainly
require an improved delivery mechanism. It may be in the
form of nanoparticles, as they are highly modular and can
carry a range of transiently expressed cargo.
122,123
By re-
administering nanoparticles to counteract the conse-
quences of muscle turnover, musculoskeletal dystrophies
could be cured.
The scarcity of published studies reveals that systemic
muscle delivery by nanoparticles remains a formidable
obstacle, most likely due to the large quantity of muscle in
the human body and the necessity to target muscles within
the thoracic cavity. Future research should prioritize the
development of an effective, muscle-specific nanoparticle
that can be delivered through the circulatory system.
The immune system poses a significant, multi-
dimensional challenge for gene therapy strategies. Pre-
existing immunity toward AAVs and Cas9 can pose a
health risk to patients. Clearing of edited cells by the im-
mune system due to the expression of Cas9 or the thera-
peutically relevant protein can reduce the effectiveness of
any strategy. The use of immunosuppressants is a potential
strategy to address the immune response elicited by
CRISPR gene therapies, particularly in the case of repeated
administrations
.124–126
However, immunosuppressants in-
crease the risk of infections, cancer, and potential organ
damage. Further work is required to investigate innate and
humoral responses to CRISPR/Cas9 gene editing strategies
in vivo and potential solutions to these challenges.
Although there are challenges to bringing therapeutic
genome editing for MDs closer to the clinic, many exciting
advances have been made to date in the field of gene-
editing in MDs. Ongoing research focusing on improving
delivery vehicles and new formulations of CRISPR com-
ponents such as mRNA and ribonucleoproteins for tran-
sient and tissue-specific delivery will undoubtedly bring
this technology to the forefront of clinical translation.
ACKNOWLEDGMENTS
The Cohn and Ivakine lab members are gratefully ac-
knowledged for their input in this work.
AUTHORS’ CONTRIBUTIONS
S.F.: Conceptualization, Data curation, Writing—
Original draft, Writing—Review and editing, Visualization.
R.M.M.: Conceptualization, Data curation, Writing—
Review and editing, Visualization.
M.J.R.: Conceptualization, Data curation, Writing—
Review and editing.
L.P.: Conceptualization, Data curation, Writing—
Review and editing.
E.A.I.: Conceptualization, Writing—Review and edit-
ing, Supervision.
R.D.C.: Conceptualization, Writing—Review and
editing, Supervision.
AUTHOR DISCLOSURE
No competing financial interests exist.
FUNDING INFORMATION
This work was funded by the Canadian Institutes of
Health Research (Ronald D. Cohn and Evgueni A. Iva-
kine), the McArthur family (Ronald D. Cohn), Jesse’s
Journey (Ronald D. Cohn), and the Michael Hyatt Foun-
dation (Ronald D. Cohn).
CRISPR/CAS9 TREATMENT OF MUSCULAR DYSTROPHIES 399
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Received for publication April 13, 2023;
accepted after revision April 24, 2023.
Published online: April 28, 2023.
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