Neuromuscular Research Centre, Departments of Neurology and Biomedicine, University Hospital Basel, Switzerland
Muscular dystrophies are classically subdivided according to their clinical phenotype, and were historically defined as progressive myopathies in which muscle biopsies demonstrate muscle fibre necrosis and regeneration, as well as replacement of muscle fibres by adipose and connective tissue. In recent years, great progress has been made in identifying the genetic basis of many myopathies, thereby presenting opportunities to develop therapeutic strategies that act on specific molecular pathomechanisms. The different therapeutic strategies and their molecular targets will be reviewed.
Key words: myopathies; dystrophinopathy; limb-girdle muscular dystrophy; dysferlinopathy; exon-skipping
The muscular dystrophies are caused by mutations in a large variety of genes with different functions, encoding proteins of the contractile apparatus, structural proteins, enzymes or nuclear proteins (fig. 1). The disease-causing mutations lead in most cases to a loss of function, but in some cases mutations can also cause a toxic gain of function. Although most genomic mutations in muscular dystrophies encode altered proteins, in some instances the genetic alterations exert their pathogenicity at the level of deoxyribonucleic acid (DNA) or ribonucleic acid (RNA). The obvious primary molecular targets in muscular dystrophy are, therefore, the affected genes and/or their respective products. Theoretically, the most effective strategy would be to correct the mutated DNA by genetic engineering. Although recent developments in genetic engineering are promising, this approach is still technically challenging and poses ethical problems. Currently, most of the therapeutic strategies are focused on the mutated gene products or their downstream targets.
Despite great efforts, a cure for muscular dystrophies is unfortunately still unavailable; however, some therapeutic strategies are currently being tested in clinical trials. The most commonly encountered forms of muscular dystrophy in adults are myotonic dystrophy (MD), facioscapulohumeral muscular dystrophy (FSHD) and the large group of limb girdle muscular dystrophies (LGMDs). The most common childhood form of muscular dystrophy is Duchenne muscular dystrophy (DMD). Experimental therapeutic efforts have therefore primarily focused on these disorders, which will be reviewed here.
Duchenne muscular dystrophy (DMD) is the most severe form of muscular dystrophy caused by mutations in the X-linked dystrophin gene. The disease affects about 1 in 3,500 live born boys, who demonstrate delayed developmental motor milestones. Gower’s manoeuvre when rising from the floor and difficulty climbing stairs in young boys raises the suspicion of DMD. Hypertrophy of the calves is present and may also affect other muscle groups. Wasting and weakness predominantly affect limb girdle muscles of the upper and lower extremities. With disease progression the child looses ambulation at around 12 years of age and becomes wheelchair-bound . Dystrophinopathy patients who remain ambulant beyond 16 years of age are considered to suffer from the Becker-type of the disease . With progression of muscle weakness, contractures develop and respiratory problems are aggravated by thoracic deformities and weakness of intercostal and diaphragmatic muscles. Cardiac problems may become apparent at a stage when muscle weakness is already pronounced.
Becker muscular dystrophy (BMD) is a milder allelic variant of DMD, with later age of onset (in very rare instances up to 60 years) and slower disease progression involving the same muscle groups as DMD, and affecting 1 in about 17,000 males . Clinical cardiac involvement is more frequent in BMD since skeletal muscle strength still allows ambulation and strain on the heart muscle is greater than in DMD [3, 4]. Cognitive dysfunction is less pronounced than in DMD patients.
X-linked dilated cardiomyopathy (XDLC) is the clinical presentation of a cardiomyopathy caused by dystrophin mutations without symptoms of skeletal muscle involvement . However, on laboratory testing serum creatine kinase levels are elevated and skeletal muscle biopsy shows myopathic changes. Typically, 20-year-old male patients present with congestive heart failure that is lethal within 1 year if not transplanted. Women may also be affected, but as a result of their mosaicism may have a slower progression of the disease.
Manifest carriers are women who carry one copy of a mutated dystrophin gene and may present with asymmetric muscle weakness or muscle hypertrophy with or without cardiomyopathy [6–9].
The dystrophin gene spans a region of about 3Mb of DNA and is composed of 79 exons . It is the largest gene known to date. The transcribed messenger RNA (mRNA) is 14kb in size and the mature protein 427kD in weight . The dystrophin molecule has an N-terminal actin-binding domain followed by 24 spectrin-like triple helical repeats that form the rod domain, which includes four nonhelical “hinge regions”. The C-terminal part contains a cysteine-rich domain that binds to beta-dystroglycan, followed by a region binding to syntrophins and dystrobrevins . Thus dystrophin provides a link between the actin cytoskeleton and the extracellular matrix (figs 1 and 2) .
The majority of mutations in dystrophin are exonic or multiexonic deletions (about 65% in DMD), less frequent are duplications (around 7%) and the remainder of the mutations are single nucleotide changes generating nonsense (DMD) or missense mutations (BMD). There are two mutational hotspots: a “major hotspot” region, which spans exons 45 to 53 and a “minor hotspot”, which spans exons 2 to 20. Mutations in DMD usually disrupt the open reading frame, whereas mutations in BMD retain the open reading frame, generating an internally truncated dystrophin molecule with sufficient biological activity to account for the milder phenotype (fig. 2) .
Several clinical trials suggest that there is a significant increase in strength, timed muscle function tests and pulmonary function in DMD patients receiving steroid treatment . Daily treatment with prednisone at a starting dosage of 0.75 mg/kg/day or deflazacort at a starting dosage of 0.9 mg/kg/day offers an effective initial treatment. This opinion is shared by a Cochrane review on glucocorticoid corticosteroid therapy in DMD  and by the report of the Quality Standards Subcommittee of the American Academy of Neurology and the Practice Committee of the Child Neurology Society . In a recent prospective longitudinal observational study, the long-term effects of intermittent and daily glucocorticoids treatment in DMD patients were compared and analysed . This study demonstrated that daily treatment was associated with longer ambulation expectancy and slower decline of motor function than the intermittent course, but the intermittent therapy was overall better tolerated with fewer adverse effects.
In recent years, however, more specific therapeutic strategies have been developed for dystrophinopathies, which include cell replacement therapies, gene therapy, the use of aminoglycosides or similar compounds allowing stop-codon read-through and the upregulation of utrophin, as well as the clinically most advanced strategy using antisense oligonucleotides for exon-skipping.
Cell replacement therapy
Experimental cell replacement therapy approaches for muscular dystrophy have been pursued for a long time and included experiments whereby normal muscle was transplanted into a dystrophic animal [19, 20]. Ethical issues, in particular nonavailability of newborn muscle that would more easily be reinnervated and revascularised, made it difficult to pursue this avenue further. A different experimental procedure consists of injecting donor muscle precursor cells (myoblasts) into a dystrophic host, where donor myoblasts, which express dystrophin, would fuse with host myotubes, originally dystrophin deficient [21, 22]. For this strategy, immune reactions against the donor cells have to be overcome [23, 24]. Other experimental strategies involve the genetic manipulation of autologous donor myoblasts ex vivo : the introduction of a dystrophin transgene restores functional dystrophin expression in injected myoblasts [26–28]. Additional cell-based experiments included systemic delivery of multipotent precursor cells, which can be isolated from nonmuscle tissue such as blood, blood vessels, bone marrow and adipose tissue . Another approach is to use induced pluripotent stem cells, which can be generated from different cell types such as fibroblasts of the skin or muscle. These pluripotent stem cells can be differentiated in vitro into muscle precursor cells before being injected systematically into the recipient host [30, 31].
Myoblasts or satellite cells were most often used in human clinical trials for dystrophinopathies. Most of these clinical trials were not successful in improving muscle strength, despite multiple injections of a large number of donor myoblasts [32–34]. The disadvantage of satellite cells is their inability to effectively cross the vessel wall and that they cannot be administered systemically but have to be injected intramuscularly.
Therefore, the systemic administration of haematopoetic stem cells (CD133+) has been evaluated in a clinical phase I trial and was shown to be safe . Dystrophin-expressing muscle fibres could be obtained in experiments applying this strategy to mdx mice (an animal model harbouring a stop codon in exon 23 of the dystrophin gene) . Experiments in animal models using mesoangioblasts demonstrated their capacity to ameliorate the phenotype in the mouse  and dog , and subsequently led to clinical trials with human allogeneic human leucocyte antigen (HLA) identical mesoangioblasts for the treatment of patients affected by DMD (https://www.clinicaltrialsregister.eu; EudraCT Nummer: 2011-000176-33), which are currently ongoing.
Gene replacement therapy
Diseases caused by deficiency of a product from a mutated gene (rather than a toxic gain of function) are, in theory, amenable to gene replacement approaches.
Experimental methods aiming at the introduction of a foreign gene into the host organism include administration of plasmid DNA  or vector-mediated gene transfer [39, 40] (fig. 3).
Several plasmid-mediated gene transfer studies have been performed in the mdx mouse, through direct injection of plasmids into skeletal muscle [41–45]. Direct plasmid DNA injection into skeletal muscle is usually less efficient than virus-mediated gene transfer, but is not associated with an immune response to the viral vector. However, the generation of immune responses to new epitopes encoded by the transgene is still a possibility . In mice, intravascular injections of dystrophin-encoding plasmids have resulted in dystrophin expression in up to 10% of muscle fibres, which lasted for 6 months . A phase I clinical trial of plasmid-mediated dystrophin complementary DNA (cDNA) delivery was conducted in Duchenne boys. A forearm muscle was injected and no adverse effects were noted. Six of the nine patients had low levels of dystrophin containing fibres after 3 weeks .
Viral vectors can enhance transgene uptake, but have other disadvantages. Commonly used viral vectors for gene transfer into muscle cells are retroviral, adenoviral, and adeno-associated viral vectors (fig. 3).
Retroviral vectors stably integrate the desired transgene into the host genome . The integration is mainly a random process and can, therefore, depending on the integration site, activate oncogenes . The development of self-inactivating lentiviral vectors, which integrate without their viral promoters and enhancing sequences, may solve this problem . The major advantage of retroviral vectors is their stable integration into the genome, resulting in a permanent expression of the transgene. This approach, in combination with cell replacement therapy, has been used to repair dystrophin deficient cells ex vivo [50, 51].
The commonly used adenoviral vectors have the advantage of a large insert capacity and the ability to infect postmitotic cells. Adenoviral vectors have been used as a therapeutic strategy for dystrophinopathies [39, 52–54]. Adenoviral vectors have an insert capacity of about 8kb. Deletion of genes from the viral genome can further increase the insert capacity of the adenovirus up to 36kb. These “gutted” adenoviruses need to be grown in the presence of helper viruses, which provide the viral gene products originally encoded by the deleted genome. Studies in mdx mice with viruses delivering dystrophin have shown restoration of the dystrophin-glycoprotein complex at the sarcolemma, long-term expression of dystrophin and correction of histological and physiological indices of muscle disease, especially in young mice [39, 55–57].
Adenovirus-associated viruses (AAVs) have a higher muscle tropism, are less immunogenic, and have not been reported to be associated with any human disease. AAV vectors, however, have a considerably smaller insert capacity of about 4kb; insertion of full length dystrophin cDNA is therefore not possible. However, small, functional “minidystrophin” molecules can be incorporated into these viral vectors and have been used in gene transfer experiments in dystrophinopathy mouse models (fig. 2) [40, 58, 59]. Stable gene expression could be observed up to 7 years after injection of AAV vectors in dogs and rhesus monkeys [60, 61]. In a human clinical trial, AAV vectors coding for minidystrophin have been injected intramuscularly into DMD patients [62, 63]. Although the viral genome could be detected in all biopsies of injected muscles, in only a few patients a small number of dystrophin-expressing muscle fibres could be identified. The observed T-cell-mediated immune response against minidystrophin in these patients could be an explanation for the low number of muscle fibres expressing dystrophin .
Mutation suppression therapy
Stop-codon read-through has been reported to be achievable with aminoglycoside antibiotics in cell culture and in vivo . As about 10% of mutations in the dystrophin gene are point mutations leading to premature termination of translation, such “read-through therapy” would be of potential therapeutic interest. Therapeutic applications could be expanded to include other muscular dystrophies or other genetic diseases arising from premature termination of translation. Mdx mice harbouring a stop codon in exon 23 treated with gentamicin showed increased dystrophin expression and partial restoration of the dystrophin glycoprotein complex (DGC), as well as protection from contraction-induced muscle fibre damage and reduction in serum creatine kinase levels . This observation, however, could not be repeated in another animal study . Clinical trials did not show dystrophin staining after 2 weeks of daily gentamicin treatment (7.5 mg/kg) in four patients . In a second study, 12 patients with either dystrophinopathy or sarcoglycanopathy were treated for 2 weeks with daily gentamicin (7.5 mg/kg) . The latter study reported a decrease of serum creatine kinase levels, but without improvement in strength, functional testing or dystrophin expression. A more recent clinical trial demonstrated that 6 months of gentamicin treatment increased dystrophin levels with reduced creatine kinase levels, suggesting that the administration of gentamicin successfully induced read-through of stop codons . However, the toxicity of gentamicin remains a major problem.
Ataluren (PTC-124, a 1,2,4-oxadiazole compound) is a small molecule that can override nonsense stop translation signals to produce full-length proteins. PTC-124 was effective in restoring the production of full-length proteins in animal models of cystic fibrosis and DMD . In a large multicentre placebo-controlled double-blind study, patients received either one of two different doses of ataluren or placebo. In a 6-minute walk test, patients with the lower dose of ataluren showed an improvement in walking distance compared with the placebo group. However, this difference was not statistically significant and the group receiving the high dose of ataluren did not show any difference compared with the placebo group [70, 71].
Exon skipping therapy
The dystrophin gene includes 79 exons, which must be spliced together to produce the mature muscle, brain and cardiac transcripts. The splicing process is made possible by the splicing machinery and specific nucleotide motifs present on the prespliced RNA that defines exon-intron boundaries. The masking of such domains does not allow exon definition and leads to exclusion of the exon with the adjacent introns from the mature messenger RNA (mRNA) transcript, thus leading to exon skipping. The dystrophin gene sequence is known and elements involved in exon definition can be predicted. Thus it is possible to design specifically antisense nucleotides that would mask such domains important for splicing and induce exon skipping. Hence, exons containing stop codons could, theoretically, be skipped with this technique. It could be envisaged that several exons could be skipped until an open reading frame is reinstalled, in the case where skipping of one particular exon leads to out-of-frame splicing. This technique could, in theory, be of use for all dystrophin mutations that do not reside in essential protein domains, which make up about 80% of mutations in Duchenne patients.
In-vitro and in-vivo experiments in the mdx mouse and in dogs have demonstrated the feasibility of the exon skipping strategy for dystrophinopathies [72–74]. Systemic delivery of antisense nucleotides (AOs) into the mdx mouse  led to low levels of dystrophin devoid of exon 23, and AOs led to expression of dystrophin in the golden retriever dog model . Exon skipping could be achieved in humans after intramuscular, as well as systemic, administration of AOs [75, 76].
In ongoing clinical trials, two different antisense molecules are being used: a 2’O-methyl phosphorothionate oligonucleotide (PRO51/GSK2402968 Prosensa Therapeutics/GSK) and a phosphorodiamidate morpholino (AVI-4658/Eteplirsen; Sarepta Therapeutics). Currently, clinical studies to demonstrate efficacy of these two molecules in DMD patients are being conducted .
Newly expressed dystrophin epitopes can trigger an immune response in a dystrophin deficient host. Upregulation of an endogenous protein with functions similar to those of dystrophin could circumvent this undesired side effect. Utrophin contains C’ and N’ terminal domains that are highly homologous to dystrophin, with less homology in the central rod domain. Utrophin preferentially accumulates at the neuromuscular and myotendinous junctions [78, 79] in intact muscle, where it has similar functions to dystrophin, whereas dystrophin is found along the entire length of the sarcolemma. Utrophin can serve as a link between the actin cytoskeleton and the extracellular matrix through association with the dystrophin glycoprotein complex . Therefore, it can be hypothesised that utrophin upregulation might correct dystrophin deficiency. Indeed, in muscle from dystrophin deficient patients, utrophin is already upregulated , possibly in an attempt by the muscle cell to compensate for dystrophin deficiency. Experimental overexpression of utrophin can lead to structural and functional improvements in mdx mice [81, 82]. Enhancement of utrophin expression through upregulation of the endogenous gene [83–85] could be of benefit without causing an immune response [86, 87].
Limb girdle muscular dystophies
The LGMDs are a heterogeneous group of muscle disorders characterised clinically by weakness and wasting in the pelvic and shoulder girdle muscles, and pathologically by muscle necrosis and regeneration. This group of disorders can be divided into two types on the basis of their pattern of inheritance: type 1 – autosomal dominant and type 2 – autosomal recessive. The dominantly inherited LGMDs are rare, representing <10% of all LGMDs. The more common autosomal recessive forms have a prevalence of about 1:15,000 to 1:100,000. LGMDs are further subdivided according to their different genetic loci (table 1).
Mutations causing LGMD are found in genes encoding proteins that are membrane-associated, cytosolic, sarcomeric or constituents of the nuclear envelope (fig. 1).
Clinically, age at onset and rate of disease progression are variable in the LGMDs, both within the same disease group and even within members of the same family. Moreover, identical mutations may cause different disease phenotypes, even among family members. These findings point to the existence of putative modifying factors. Such phenotypic variability makes prediction about disease course difficult and should be considered when counselling patients. Currently, immunohistochemistry of histological sections and Western blot analysis on muscle homogenates are performed as diagnostic steps to determine the putative protein defects in LGMD. However, in many instances, reduction of a given protein may be secondary to the absence of another. This points to shared pathophysiological pathways, but also poses diagnostic difficulties. Therefore, whenever possible, mutational analysis should be performed to confirm the biochemical findings.
|Table 1: Limb girdle muscular dystrophies (LGMDs).|
AD = autosomal dominant
AR = autosomal recessive
|LGMD 1B||AD||Lamin A/C||1q22|||
|LGMD 2C||AR||Sarcoglycan, gamma||13q12|||
|LGMD 2D||AR||Sarcoglycan, alpha||17q21|||
|LGMD 2E||AR||Sarcoglycan, beta||4q12|||
|LGMD 2F||AR||Sarcoglycan, delta||5q33-q34|||
|LGMD 2L||AR||Anoctamin 5||11p14-12|||
|LGMD 2Q||AR||Plectin 1||8q24|||
|DNAJB6 = DnaJ (Hsp40) homolog, subfamily B, member 6; FKRP = fukutin-related protein; POMGNT1 = protein O-linked mannose N-acetylglucosaminyltransferase 1; POMT1 = protein O-mannosyl transferase 1; POMT2 = protein O-mannosyl transferase 2; TRIM32 = tripartite motif containing 32|
Dysferlin is a membrane protein of about 230kD that contains several C2 motifs. These motifs are present in many membrane-associated proteins, and bind calcium and anionic phospholipids [88–90]. Dysferlin is necessary for the repair of membrane tears in muscle cells, which can arise during muscle exercise . Loss of dysferlin leads to a defective repair mechanism and the integrity of the plasma membrane cannot be restored following injury, leading to muscle fibre necrosis (fig. 4B).
Dysferlin mutations can result in two distinct phenotypes that are characterised by either proximal or distal weakness and wasting; the distal form is known as Miyoshi myopathy and the proximal form as LGMD2B. Both phenotypes can be present within the same family [92, 93]. Dysferlin mutations account for about 20% of patients with recessively inherited LGMDs. Facial and pharyngeal muscles remain unaffected. Cardiac, respiratory or cognitive impairment is not part of the disease, although some reports mention cardiac involvement . Serum creatine kinase levels can be very high (up to 150 times normal) and, occasionally, inflammatory infiltrates are seen on skeletal muscle biopsy, which may be mistaken for a primary inflammatory myopathy. Age of onset can be variable, but is mostly in the teenage years. Patients can be engaged in athletic activities prior to disease onset.
Several therapeutic strategies similar to the approaches described for DMD have been applied in experiments: exon skipping strategies [23, 95, 96], stop codon read through  and somatic gene therapy using minidysferlins [98, 99]. Another strategy is the inhibition of degradation of missense-mutated dysferlin [100, 101].
The elimination of mutated and misfolded proteins is conducted by the cellular quality control system, which recognises misfolded proteins and initiates their degradation to prevent aggregation within the cell. In-vitro experiments on muscle cells from dysferlinopathy patients demonstrated that missense mutations destabilise the dysferlin protein and lead to its rapid degradation. Treatment of patient-derived myoblasts with inhibitors of the proteasome prevented the degradation of the mutated protein and restored the membrane repair function of the cultured muscle cells. Currently, there is an ongoing clinical trial to validate this strategy in dysferlinopathy patients (NCT01863004 / clinicaltrials.gov)
Myotonic dystrophy is an autosomal dominant multisystem disease affecting skeletal muscle, heart, brain, lens and endocrine organs. On the basis of genetic loci and clinical characteristics, two myotonic dystrophies can be distinguished: type 1 (DM1, Curschmann-Steinert) and type 2 (proximal myotonic myopathy, PROMM, DM2). DM1 is caused by a CTG expansion in the 3’ untranslated region of the DMPK gene , and DM2 by a CCTG expansion in intron 1 of the ZNF9 gene .
The clinical findings in DM1 have been subdivided according to severity into three overlapping phenotypes: mild, classic and congenital. These phenotypes roughly correlate with CTG expansion size. Individuals with the mild form of DM1 usually have a CTG repeat size between 50 and 150, age at onset is between 20 and 60 years, life-span is unaffected and disease may manifest itself with cataracts, diabetes and mild myotonia.
Patients affected by the classical form of DM1 have a CTG repeat size of up to 1,000 and develop muscle weakness and wasting, myotonia, cataracts, diabetes and heart conduction abnormalities. A myopathic facial expression can develop as a result of weakness of the facial muscles. Muscle weakness predominantly affects distal muscles in the lower and upper extremities. Smooth muscle involvement can result in swallowing difficulties and difficulties with bowel movements.
Life-span is mildly reduced and the most common cause of death is respiratory insufficiency due to diaphragmatic, intercostal and orpharyngeal involvement . Cardiac sudden death caused by heart block and ventricular arrhythmia is the second most common cause of death in DM1 patients.
Central nervous system manifestations of the disease may include hypersomnia, personality disorders, altered regulation of respiration and cognitive impairment. Other manifestations of the disease include cataracts of the posterior subcapsular zone, testicular atrophy, balding, and diabetes due to insulin resistance. Women affected by DM1 may present problems during pregnancy and labour, such as prolonged labour, retained placenta and postpartum haemorrhage, owing to uterine smooth muscle involvement. The spontaneous abortion rate is higher in women with DM1, and if the foetus is affected by a congenital form of DM1, further complications include polyhydramnions and reduced foetal movements.
Children with the congenital form of DM1 have usually over 2,000 CTG repeats and they may present prenatally with polyhydramnions. Neonatal hyptonia, feeding difficulty and respiratory compromise may be present . Ensuing from facial muscle weakness in utero is the typical V-shaped (tented) mouth. Mental retardation is present in about half of the patients with congenital DM1 and some patients have a certain degree of cerebral atrophy and ventricular dilatation. About a quarter of children with DM1 die from failure to develop adequate respiratory function, and surviving patients have a reduced life-span .
Current treatment is symptomatic, and involves vigilant cardiac monitoring and implantation of a pacemaker when indicated, cataract removal, and treatment of diabetes and of pain. Adaptation of assistive devices as disease progresses is helpful. Daytime sleepiness can be treated with modafinil [107, 108].
DM1 is caused by an unstable CTG repeat expansion in the 3’ untranslated region of the myotonic dystrophy protein kinase gene (DMPK) (fig. 4A) . As the unstable repeats enlarge when transmitted from one generation to the next, subsequent generations are expected to be more affected than their parents. Repeats smaller than 100 tend to be more unstable in sperm than in ova  and the transition from mild to classical DM1 is usually transmitted by the father. However, congenital DM1 is usually transmitted by the mother, suggesting that sperm probably are not functional when harbouring very large repeats .
Apart from germline instability, there is also somatic instability, with over a 10-fold increase of repeat length in skeletal muscle compared with DNA derived from blood leucocytes. Prolongation of nucleotide repeats probably occurs during DNA repair .
The triplet repeat in DM1 is located in the 3’ untranslated region of the DMPK gene. Although many studies suggest that DMPK levels are reduced in DM1 , DMPK-deficient mice show only a subtle phenotype with first degree atrioventricular (AV) cardiac conduction block without myotonia in the heterozygous state [113, 114]. These experiments suggest that DMPK is dispensable for muscle development and that a reduction of DMPK may account at most for only a few of the features seen in DM1. The finding that an intronic nucleotide repeat expansion on a different chromosome causes DM2 with very similar features to DM1 further questions whether haploinsufficiency of DMPK can account for the disease phenotype of DM.
Transgenic mice with expanded CTG repeats introduced into the 3’ untranslated region of a human skeletal actin gene develop myotonia and muscle pathology similar to patients with DM1 . Transgenic mRNA accumulates in the nucleus and forms aggregates similar to DMPK mRNA in DM1 patients. These elongated CUG repeats form hairpin structures and can sequester RNA-binding proteins implicated in splicing, such as the protein muscleblind-like-1 (MBNL1) [116-118]. Human muscleblind-like genes MBNL1, MBNL2, and MBNL3 are homologous with the Drosophila gene muscleblind, which is essential for muscle and eye differentiation .
Muscleblind is implicated in splicing the chloride channel CLCN-1, which is developmentally regulated. Mutations in CLCN-1 can lead to the dominantly (Thompson) and recessively (Becker) inherited myotonias. A broad range of splicing alterations in CLCN-1 was observed in muscle tissue from DM1 and DM2 patients . Thus, alteration of CLCN-1 splicing can explain myotonia in DM. Along these lines, MBNL1 deficient mice show reduced expression of CLCN-1 owing to aberrant splicing, and exhibit myotonia. Furthermore, MBNL1 knock-out mice recapitulate many features of the DM phenotype . Other genes are incorrectly spliced in DM, such as the insulin receptor, leading to expression of the nonmuscle insulin receptor in muscle tissue. The nonmuscle insulin receptor has a decreased metabolic response to insulin. MBNL deficient mice also develop cataracts . The CUGBP/Elav like family member 1 (CELF1) is another splicing factor, which is misregulated owing to the trinucleotide expansions and seems to be involved in the missplicing phenotype. CELF1 levels are elevated and the protein is hyperphosphorylated, leading to a missplicing of certain gene transcripts, in DM1 [122, 123].
The concept of a “splice-opathy” as the pathogenic mechanism raises very interesting potential therapeutic possibilities. First, muscle damage is not well advanced early in the disease in DM and is only slowly progressive, suggesting that restoration of the splicing abnormality may allow “recovery” of muscle tissue. The hairpin loops formed by expanded triplet repeats that bind and sequester MBLN proteins could be targeted by low molecular weight compounds with the idea of disrupting the hairpin structure, and thus interfering with MBLN binding (fig. 4A). Different therapeutic strategies have been tested in in-vitro or in-vivo experiments using animal models: antisense technologies [124–127], RNA interference [128, 129] and ribozyme technology , as well as peptides  and small molecular compounds [132–134].
Facioscapulohumeral muscular dystrophy
Facioscapulohumeral muscular dystrophy (FSHD, MIM 158900) is an autosomal dominantly inherited neuromuscular disorder with an incidence of approximately 1 in 20,000 . Typical age at diagnosis is in the second decade of life. Symptoms can range from severe upper and lower limb involvement and wheelchair use in early childhood, to very mild facial weakness in the seventh decade . Scapula-fixators are usually first affected; rarely the disease begins with weakness of facial, foot-dorsiflexor or pelvic girdle muscles. Shoulder girdle weakness leads to asymmetrical scapula alata and pectoralis muscle atrophy. An initially fairly normal deltoid muscle contributes to the distinctive high rise of the scapula on elevation of the arms. Another very typical feature is the asymmetrical facial weakness. Muscle pain and fatigue are frequently reported; dysphagia is rare, as is respiratory insufficiency [137, 138]. Occasionally, cardiac conduction defects have been observed, although most authors claim no cardiac involvement. A subclinical high-tone hearing loss and a subclinical retinal vasculopathy have been described as part of the disease [139, 140].
The FSHD locus maps to chromosome 4q35 adjacent to the 4qter subtelomeric region . A variable number of tandem repeats, designated D4Z4, has been identified in this region, which in normal individuals usually contains multiple integral copies (11‒>100) of a 3.3kb tandem repeat sequence . The number of the repeats is reduced in the vast majority of FSHD patients to less than 11.
Recent studies have uncovered the pathomechanism of FSHD. Experiments demonstrated that induced expression of DUX4, a transcriptional factor within the DZ4Z repeats, initiates a gene expression programme toxic for muscle cells. The DUX4 gene is normally epigenetically repressed. There are two requirements for developing the disease: (1.) induction of DUX4 transcription by epigenetic activation and (2.) stabilisation of the DUX4 mRNA transcript allowing translation of the DUX4 protein (fig. 4C). In FSHD1, the epigenetic activation is induced by the reduction of the DZ4Z repeat number to below 11. The deletions lead to hypomethylation and finally transcription of the DUX4 gene locus. FSHD2 patients have normal DZ4Z repeat numbers, but show hypomethylation and active transcription of the DUX4 gene. A recent study showed that in a subset of FSHD2 patients the epigenetic activation is a consequence of a mutation in the SMCHD1 gene.
A second genetic variant is necessary to allow translation of the DUX4 protein: Two distinct 4qter subtelomeres, designated 4qA and 4qB, have been recognised [143, 144]. The 4qA- and 4qB-defined 4qter subtelomeres are found to occur with almost equal frequency in the population . However, in essentially all FSHD patients, disease expression is not only associated with large D4Z4 contractions, but these contractions must be specifically located on a 4qA-defined 4qter subtelomere [143, 144]. The 4qA variant has a polyadenylation site stabilising the DUX4 mRNA and allowing translation of DUX4 protein. Thus, induction of DUX4 transcription by epigenetic activation and stable DUX4 mRNA transcripts need to be present for disease development (fig. 4C).
Possible therapeutic strategies could target the DUX4 mRNA for degradation to prevent DUX4 protein production within muscle cells or directly target the DUX4 protein to inhibit its function as a transcription factor. Recent insights into the molecular pathomechanism of FSHD will allow the development of specific drugs targeting either DUX4 mRNA or protein.
Funding / potential competing interests: No financial support and no other potential conflict of interest relevant to this article was reported. The Neuromuscular Research group is supported by grants from the SNF, Gerbert-Rüf, Schweizer Stiftung zur Erforschung der Muskelkrankheiten und Schweizerische Muskelgesellschaft.
Correspondence: Professor Michael Sinnreich, MD, PhD, Neuromuscular Research Centre, Departments of Neurology and Biomedicine, University Hospital Basel, Petersgraben 4, CH-4031 Basel, Switzerland, Michael.sinnreich[at]unibas.ch
1 Emery AE. Duchenne Muscular Dystrophy. New York: Oxford University Press; 1993.
2 Emery AE, Skinner R. Clinical studies in benign (Becker type) X-linked muscular dystrophy. Clin Genet. 1976;10(4):189–201. PubMed PMID: 975594.
3 Nigro G, Comi LI, Politano L, Bain RJ. The incidence and evolution of cardiomyopathy in Duchenne muscular dystrophy. Int J Cardiol. 1990;26(3):271–7. PubMed PMID: 2312196..
4 Nigro G, Comi LI, Politano L, Limongelli FM, Nigro V, De Rimini ML, et al. Evaluation of the cardiomyopathy in Becker muscular dystrophy. Muscle Nerve. 1995;18(3):283–91. PubMed PMID: 7870105.
5 Mestroni L, Giacca M. Molecular genetics of dilated cardiomyopathy. Curr Opin Cardiol. 1997;12(3):303–9. PubMed PMID: 9243088.
6 Grain L, Cortina-Borja M, Forfar C, Hilton-Jones D, Hopkin J, Burch M. Cardiac abnormalities and skeletal muscle weakness in carriers of Duchenne and Becker muscular dystrophies and controls. Neuromuscul Disord. 2001;11(2):186–91. PubMed PMID: 11257476.
7 Mercier S, Toutain A, Toussaint A, Raynaud M, de Barace C, Marcorelles P, et al. Genetic and clinical specificity of 26 symptomatic carriers for dystrophinopathies at pediatric age. Eur J Hum Genet. 2013;21(8):855–63. PubMed PMID: 23299919. Pubmed Central PMCID: PMC3722679.
8 Viggiano E, Picillo E, Cirillo A, Politano L. Comparison of X-chromosome inactivation in Duchenne muscle/myocardium-manifesting carriers, non-manifesting carriers and related daughters. Clin Genet. 2013;84(3):265–70. PubMed PMID: 23110537.
9 Soltanzadeh P, Friez MJ, Dunn D, von Niederhausern A, Gurvich OL, Swoboda KJ, et al. Clinical and genetic characterization of manifesting carriers of DMD mutations. Neuromuscul Disord. 2010;20(8):499–504. PubMed PMID: 20630757. Pubmed Central PMCID: PMC2944769.
10 Nobile C, Marchi J, Nigro V, Roberts RG, Danieli GA. Exon-intron organization of the human dystrophin gene. Genomics. 1997;45(2):421–4. PubMed PMID: 9344670.
11 Hoffman EP, Brown RH, Jr., Kunkel LM. Dystrophin: the protein product of the Duchenne muscular dystrophy locus. Cell. 1987;51(6):919–28. PubMed PMID: 3319190.
12 Culligan KG, Mackey AJ, Finn DM, Maguire PB, Ohlendieck K. Role of dystrophin isoforms and associated proteins in muscular dystrophy (review). Int J Mol Med. 1998;2(6):639–48. PubMed PMID: 9850730.
13 Ervasti JM, Campbell KP. Membrane organization of the dystrophin-glycoprotein complex. Cell. 1991;66(6):1121–31. PubMed PMID: 1913804.
14 Aartsma-Rus A, Van Deutekom JC, Fokkema IF, Van Ommen GJ, Den Dunnen JT. Entries in the Leiden Duchenne muscular dystrophy mutation database: an overview of mutation types and paradoxical cases that confirm the reading-frame rule. Muscle Nerve. 2006;34(2):135–44. PubMed PMID: 16770791.
15 Manzur AY, Kuntzer T, Pike M, Swan A. Glucocorticoid corticosteroids for Duchenne muscular dystrophy. Cochrane Database Syst Rev. 2008(1). PubMed PMID: WOS:000252926800138.
16 Manzur AY, Kuntzer T, Pike M, Swan A. Glucocorticoid corticosteroids for Duchenne muscular dystrophy. Cochrane Database Syst Rev. 2004(2):CD003725. PubMed PMID: 15106215.
17 Moxley RT, 3rd, Ashwal S, Pandya S, Connolly A, Florence J, Mathews K, et al. Practice parameter: corticosteroid treatment of Duchenne dystrophy: report of the Quality Standards Subcommittee of the American Academy of Neurology and the Practice Committee of the Child Neurology Society. Neurology. 2005;64(1):13–20. PubMed PMID: 15642897.
18 Ricotti V, Ridout DA, Scott E, Quinlivan R, Robb SA, Manzur AY, et al. Long-term benefits and adverse effects of intermittent versus daily glucocorticoids in boys with Duchenne muscular dystrophy. J Neurol Neurosurg Psychiatry. 2013;84(6):698–705. PubMed PMID: 23250964.
19 Partridge TA, Grounds M, Sloper JC. Evidence of fusion between host and donor myoblasts in skeletal muscle grafts. Nature. 1978;273(5660):306–8. PubMed PMID: 652035.
20 Law PK, Yap JL. New muscle transplant method produces normal twitch tension in dystrophic muscle. Muscle Nerve. 1979;2(5):356–63. PubMed PMID: 492212.
21 Skuk D, Goulet M, Roy B, Chapdelaine P, Bouchard JP, Roy R, et al. Dystrophin expression in muscles of duchenne muscular dystrophy patients after high-density injections of normal myogenic cells. J Neuropathol Exp Neurol. 2006;65(4):371–86. PubMed PMID: 16691118.
22 Skuk D, Goulet M, Roy B, Piette V, Côté CH, Chapdelaine P, et al. First test of a “high-density injection” protocol for myogenic cell transplantation throughout large volumes of muscles in a Duchenne muscular dystrophy patient: eighteen months follow-up. Neuromuscul Disord. 2007;17(1):38–46. PubMed PMID: 17142039.
23 Wein N, Avril A, Bartoli M, Beley C, Chaouch S, Laforêt P, et al. Efficient bypass of mutations in dysferlin deficient patient cells by antisense-induced exon skipping. Hum Mutat. 2010;31(2):136–42. PubMed PMID: 19953532.
24 Skuk D, Paradis M, Goulet M, Tremblay JP. Ischemic central necrosis in pockets of transplanted myoblasts in nonhuman primates: implications for cell-transplantation strategies. Transplantation. 2007;84(10):1307–15. PubMed PMID: 18049116.
25 Torrente Y, Belicchi M, Marchesi C, D'Antona G, Cogiamanian F, Pisati F, et al. Autologous transplantation of muscle-derived CD133(+) stem cells in Duchenne muscle patients. Cell Transplant. 2007;16(6):563–77. PubMed PMID: WOS:000249513200001.
26 Moisset PA, Skuk D, Asselin I, Goulet M, Roy B, Karpati G, et al. Successful transplantation of genetically corrected DMD myoblasts following ex vivo transduction with the dystrophin minigene. Biochem Biophys Res Commun. 1998;247(1):94–9. PubMed PMID: 9636661.
27 Moisset PA, Gagnon Y, Karpati G, Tremblay JP. Expression of human dystrophin following the transplantation of genetically modified mdx myoblasts. Gene Ther. 1998;5(10):1340–6. PubMed PMID: 9930339.
28 Ikemoto M, Fukada S, Uezumi A, Masuda S, Miyoshi H, Yamamoto H, et al. Autologous transplantation of SM/C-2.6(+) satellite cells transduced with micro-dystrophin CS1 cDNA by lentiviral vector into mdx mice. Mol Ther. 2007;15(12):2178–85. PubMed PMID: 17726457.
29 Tedesco FS, Dellavalle A, Diaz-Manera J, Messina G, Cossu G. Repairing skeletal muscle: regenerative potential of skeletal muscle stem cells. J Clin Invest. 2010;120(1):11–9. PubMed PMID: 20051632. Pubmed Central PMCID: PMC2798695..
30 Kazuki Y, Hiratsuka M, Takiguchi M, Osaki M, Kajitani N, Hoshiya H, et al. Complete genetic correction of ips cells from Duchenne muscular dystrophy. Mol Ther. 2010;18(2):386–93. PubMed PMID: 19997091. Pubmed Central PMCID: PMC2839293..
31 Goudenege S, Lebel C, Huot NB, Dufour C, Fujii I, Gekas J, et al. Myoblasts derived from normal hESCs and dystrophic hiPSCs efficiently fuse with existing muscle fibers following transplantation. Mol Ther. 2012;20(11):2153–67. PubMed PMID: 22990676. Pubmed Central PMCID: PMC3498803..
32 Karpati G, Carpenter S, Morris GE, Davies KE, Guerin C, Holland P. Localization and quantitation of the chromosome 6-encoded dystrophin-related protein in normal and pathological human muscle. J Neuropathol Exp Neurol. 1993;52(2):119–28. PubMed PMID: 8440993.
33 Gussoni E, Pavlath GK, Lanctot AM, Sharma KR, Miller RG, Steinman L, et al. Normal dystrophin transcripts detected in Duchenne muscular dystrophy patients after myoblast transplantation. Nature. 1992;356(6368):435–8. PubMed PMID: 1557125.
34 Mendell JR, Kissel JT, Amato AA, King W, Signore L, Prior TW, et al. Myoblast transfer in the treatment of Duchenne’s muscular dystrophy. N Engl J Med. 1995;333(13):832–8. PubMed PMID: 7651473.
35 Torrente Y, Belicchi M, Sampaolesi M, Pisati F, Meregalli M, D’Antona G, et al. Human circulating AC133(+) stem cells restore dystrophin expression and ameliorate function in dystrophic skeletal muscle. J Clin Invest. 2004;114(2):182–95. PubMed PMID: 15254585. Pubmed Central PMCID: PMC449743.
36 Sampaolesi M, Torrente Y, Innocenzi A, Tonlorenzi R, D'Antona G, Pellegrino MA, et al. Cell therapy of alpha-sarcoglycan null dystrophic mice through intra-arterial delivery of mesoangioblasts. Science. 2003;301(5632):487–92. PubMed PMID: WOS:000184340500034.
37 Sampaolesi M, Blot S, D’Antona G, Granger N, Tonlorenzi R, Innocenzi A, et al. Mesoangioblast stem cells ameliorate muscle function in dystrophic dogs. Nature. 2006;444(7119):574–9. PubMed PMID: 17108972.
38 Wolff JA, Malone RW, Williams P, Chong W, Acsadi G, Jani A, et al. Direct gene transfer into mouse muscle in vivo. Science. 1990;247(4949 Pt 1):1465–8. PubMed PMID: 1690918.
39 DelloRusso C, Scott JM, Hartigan-O’Connor D, Salvatori G, Barjot C, Robinson AS, et al. Functional correction of adult mdx mouse muscle using gutted adenoviral vectors expressing full-length dystrophin. Proc Natl Acad Sci U S A. 2002;99(20):12979–84. PubMed PMID: 12271128.
40 Watchko J, O’Day T, Wang B, Zhou L, Tang Y, Li J, et al. Adeno-associated virus vector-mediated minidystrophin gene therapy improves dystrophic muscle contractile function in mdx mice. Hum Gene Ther. 2002;13(12):1451–60. PubMed PMID: 12215266.
41 Wolff JA, Ludtke JJ, Acsadi G, Williams P, Jani A. Long-term persistence of plasmid DNA and foreign gene expression in mouse muscle. Hum Mol Genet. 1992;1(6):363–9. PubMed PMID: 1301910.
42 Acsadi G, Dickson G, Love DR, Jani A, Walsh FS, Gurusinghe A, et al. Human dystrophin expression in mdx mice after intramuscular injection of DNA constructs. Nature. 1991;352(6338):815–8. PubMed PMID: 1881437.
43 Liu F, Nishikawa M, Clemens PR, Huang L. Transfer of full-length Dmd to the diaphragm muscle of Dmd(mdx/mdx) mice through systemic administration of plasmid DNA. Mol Ther. 2001;4(1):45–51. PubMed PMID: 11472105.
44 Liang KW, Nishikawa M, Liu F, Sun B, Ye Q, Huang L. Restoration of dystrophin expression in mdx mice by intravascular injection of naked DNA containing full-length dystrophin cDNA. Gene Ther. 2004;11(11):901–8. PubMed PMID: 14985786.
45 Zhang G, Ludtke JJ, Thioudellet C, Kleinpeter P, Antoniou M, Herweijer H, et al. Intraarterial delivery of naked plasmid DNA expressing full-length mouse dystrophin in the mdx mouse model of duchenne muscular dystrophy. Hum Gene Ther. 2004;15(8):770–82. PubMed PMID: 15319034.
46 Romero NB, Braun S, Benveniste O, Leturcq F, Hogrel JY, Morris GE, et al. Phase I study of dystrophin plasmid-based gene therapy in Duchenne/Becker muscular dystrophy. Hum Gene Ther. 2004;15(11):1065–76. PubMed PMID: 15610607.
47 Li S, Kimura E, Fall BM, Reyes M, Angello JC, Welikson R, et al. Stable transduction of myogenic cells with lentiviral vectors expressing a minidystrophin. Gene Ther. 2005;12(14):1099–108. PubMed PMID: WOS:000230323100002.
48 Hacein-Bey-Abina S, von Kalle C, Schmidt M, Le Deist F, Wulffraat N, McIntyre E, et al. A serious adverse event after successful gene therapy for X-linked severe combined immunodeficiency. N Engl J Med. 2003;348(3):255–6. PubMed PMID: WOS:000180390600013.
49 Yi Y, Hahm SH, Lee KH. Retroviral gene therapy: safety issues and possible solutions. Curr Gene Ther. 2005;5(1):25–35. PubMed PMID: 15638709.
50 Bachrach E, Perez AL, Choi YH, Illigens BMW, Jun SJ, Del Nido P, et al. Muscle engraftment of myogenic progenitor cells following intraarterial transplantation. Muscle Nerve. 2006;34(1):44–52. PubMed PMID: WOS:000238688000004.
51 Dellavalle A, Sampaolesi M, Tonlorenzi R, Tagliafico E, Sacchetti B, Perani L, et al. Pericytes of human skeletal muscle are myogenic precursors distinct from satellite cells. Nature Cell Biol. 2007;9(3):255–U30. PubMed PMID: WOS:000244558600010.
52 Acsadi G, Lochmuller H, Jani A, Huard J, Massie B, Prescott S, et al. Dystrophin expression in muscles of mdx mice after adenovirus-mediated in vivo gene transfer. Hum Gene Ther. 1996;7(2):129–40. PubMed PMID: WOS:A1996UG13500001.
53 Vincent N, Ragot T, Gilgenkrantz H, Couton D, Chafey P, Gregoire A, et al. Long-term correction of mouse dystrophic degeneration by adenovirus-mediated transfer of a minidystrophin gene. Nature Genet. 1993;5(2):130–4. PubMed PMID: WOS:A1993MA05600009.
54 Clemens PR, Kochanek S, Sunada Y, Chan S, Chen HH, Campbell KP, et al. In vivo muscle gene transfer of full-length dystrophin with an adenoviral vector that lacks all viral genes. Gene Ther. 1996;3(11):965–72. PubMed PMID: WOS:A1996VR64300004.
55 Gilbert R, Dudley RW, Liu AB, Petrof BJ, Nalbantoglu J, Karpati G. Prolonged dystrophin expression and functional correction of mdx mouse muscle following gene transfer with a helper-dependent (gutted) adenovirus-encoding murine dystrophin. Hum Mol Genet. 2003;12(11):1287–99. PubMed PMID: 12761044.
56 Dudley RW, Lu Y, Gilbert R, Matecki S, Nalbantoglu J, Petrof BJ, et al. Sustained improvement of muscle function one year after full-length dystrophin gene transfer into mdx mice by a gutted helper-dependent adenoviral vector. Hum Gene Ther. 2004;15(2):145–56. PubMed PMID: 14975187.
57 Scott JM, Li S, Harper SQ, Welikson R, Bourque D, DelloRusso C, et al. Viral vectors for gene transfer of micro-, mini-, or full-length dystrophin. Neuromuscul Disord. 2002;12(Suppl 1):S23–9. PubMed PMID: 12206791.
58 Wang B, Li J, Xiao X. Adeno-associated virus vector carrying human minidystrophin genes effectively ameliorates muscular dystrophy in mdx mouse model. Proc Natl Acad Sci U S A. 2000;97(25):13714–9. PubMed PMID: 11095710.
59 Gregorevic P, Blankinship MJ, Allen JM, Crawford RW, Meuse L, Miller DG, et al. Systemic delivery of genes to striated muscles using adeno-associated viral vectors. Nat Med. 2004;10(8):828–34. PubMed PMID: 15273747.
60 Herzog RW, Yang EY, Couto LB, Hagstrom JN, Elwell D, Fields PA, et al. Long-term correction of canine hemophilia B by gene transfer of blood coagulation factor IX mediated by adeno-associated viral vector. Nat Med. 1999;5(1):56–63. PubMed PMID: WOS:000077885000030.
61 Manno CS, Chew AJ, Hutchison S, Larson PJ, Herzog RW, Arruda VP, et al. AAV-mediated factor IX gene transfer to skeletal muscle in patients with severe hemophilia B. Blood. 2003;101(8):2963–72. PubMed PMID: WOS:000182101400015.
62 Mendell JR, Campbell K, Rodino-Klapac L, Sahenk Z, Shilling C, Lewis S, et al. Dystrophin immunity in Duchenne’s muscular dystrophy. N Engl J Med. 2010;363(15):1429–37. PubMed PMID: 20925545. Pubmed Central PMCID: PMC3014106.
63 Bowles DE, McPhee SW, Li C, Gray SJ, Samulski JJ, Camp AS, et al. Phase 1 gene therapy for Duchenne muscular dystrophy using a translational optimized AAV vector. Mol Ther. 2012;20(2):443–55. PubMed PMID: 22068425. Pubmed Central PMCID: PMC3277234.
64 Barton-Davis ER, Cordier L, Shoturma DI, Leland SE, Sweeney HL. Aminoglycoside antibiotics restore dystrophin function to skeletal muscles of mdx mice. J Clin Invest. 1999;104(4):375–81. PubMed PMID: 10449429.
65 Dunant P, Walter MC, Karpati G, Lochmuller H. Gentamicin fails to increase dystrophin expression in dystrophin-deficient muscle. Muscle Nerve. 2003;27(5):624–7. PubMed PMID: 12707984.
66 Wagner KR, Hamed S, Hadley DW, Gropman AL, Burstein AH, Escolar DM, et al. Gentamicin treatment of Duchenne and Becker muscular dystrophy due to nonsense mutations. Ann Neurol. 2001;49(6):706–11. PubMed PMID: 11409421.
67 Serrano C WC, Moore. Gentamicin treatment for muscular dystrophy with stop codon mutations. Neurology. 2001;56(Suppl 3): A79.
68 Malik V, Rodino-Klapac LR, Viollet L, Wall C, King W, Al-Dahhak R, et al. Gentamicin-induced readthrough of stop codons in Duchenne muscular dystrophy. Ann Neurol. 2010;67(6):771–80. PubMed PMID: 20517938.
69 Aurino S, Nigro V. Readthrough strategies for stop codons in Duchenne muscular dystrophy. Acta Myol. 2006;25(1):5–12. PubMed PMID: 17039975.
70 Finkel RS. Read-through strategies for suppression of nonsense mutations in Duchenne/ Becker muscular dystrophy: aminoglycosides and ataluren (PTC124). J Child Neurol. 2010;25(9):1158–64. PubMed PMID: 20519671.
71 Hoffman EP, Bronson A, Levin AA, Takeda S, Yokota T, Baudy AR, et al. Restoring dystrophin expression in duchenne muscular dystrophy muscle progress in exon skipping and stop codon read through. Am J Pathol. 2011;179(1):12-22. PubMed PMID: 21703390. Pubmed Central PMCID: PMC3124804.
72 Dunckley MG, Manoharan M, Villiet P, Eperon IC, Dickson G. Modification of splicing in the dystrophin gene in cultured Mdx muscle cells by antisense oligoribonucleotides. Hum Mol Genet. 1998;7(7):1083–90. PubMed PMID: 9618164.
73 Lu QL, Rabinowitz A, Chen YC, Yokota T, Yin H, Alter J, et al. Systemic delivery of antisense oligoribonucleotide restores dystrophin expression in body-wide skeletal muscles. Proc Natl Acad Sci U S A. 2005;102(1):198–203. PubMed PMID: 15608067.
74 McClorey G, Moulton HM, Iversen PL, Fletcher S, Wilton SD. Antisense oligonucleotide-induced exon skipping restores dystrophin expression in vitro in a canine model of DMD. Gene Ther. 2006;13(19):1373–81. PubMed PMID: 16724091.
75 Cirak S, Arechavala-Gomeza V, Guglieri M, Feng L, Torelli S, Anthony K, et al. Exon skipping and dystrophin restoration in patients with Duchenne muscular dystrophy after systemic phosphorodiamidate morpholino oligomer treatment: an open-label, phase 2, dose-escalation study. Lancet. 2011;378(9791):595–605. PubMed PMID: 21784508. Pubmed Central PMCID: PMC3156980.
76 Goemans NM, Tulinius M, van den Akker JT, Burm BE, Ekhart PF, Heuvelmans N, et al. Systemic Administration of PRO051 in Duchenne's Muscular Dystrophy. N Engl J Med. 2011;364(16):1513–22. PubMed PMID: WOS:000289722700008.
77 Opar A. Exon-skipping drug pulls ahead in muscular dystrophy field. Nat Med. 2012;18(9):1314. PubMed PMID: 22961147.
78 Ohlendieck K, Ervasti JM, Matsumura K, Kahl SD, Leveille CJ, Campbell KP. Dystrophin-related protein is localized to neuromuscular junctions of adult skeletal muscle. Neuron. 1991;7(3):499–508. PubMed PMID: 1654951.
79 Khurana TS, Watkins SC, Chafey P, Chelly J, Tome FM, Fardeau M, et al. Immunolocalization and developmental expression of dystrophin related protein in skeletal muscle. Neuromuscul Disord. 1991;1(3):185–94. PubMed PMID: 1822793.
80 Matsumura K, Ervasti JM, Ohlendieck K, Kahl SD, Campbell KP. Association of dystrophin-related protein with dystrophin-associated proteins in mdx mouse muscle. Nature. 1992;360(6404):588–91. PubMed PMID: 1461282.
81 Tinsley JM, Blake DJ, Roche A, Fairbrother U, Riss J, Byth BC, et al. Primary structure of dystrophin-related protein. Nature. 1992;360(6404):591–3. PubMed PMID: 1461283.
82 Tinsley J, Deconinck N, Fisher R, Kahn D, Phelps S, Gillis JM, et al. Expression of full-length utrophin prevents muscular dystrophy in mdx mice. Nat Med. 1998;4(12):1441–4. PubMed PMID: 9846586.
83 Moorwood C, Lozynska O, Suri N, Napper AD, Diamond SL, Khurana TS. Drug discovery for Duchenne muscular dystrophy via utrophin promoter activation screening. PLoS One. 2011;6(10):e26169. PubMed PMID: 22028826. Pubmed Central PMCID: PMC3197614.
84 Tinsley JM, Fairclough RJ, Storer R, Wilkes FJ, Potter AC, Squire SE, et al. Daily Treatment with SMTC1100, a Novel Small Molecule Utrophin Upregulator, Dramatically Reduces the Dystrophic Symptoms in the mdx Mouse. Plos One. 2011;6(5). PubMed PMID: WOS:000290305600014.
85 Krag TO, Bogdanovich S, Jensen CJ, Fischer MD, Hansen-Schwartz J, Javazon EH, et al. Heregulin ameliorates the dystrophic phenotype in mdx mice. Proc Natl Acad Sci U S A. 2004;101(38):13856–60. PubMed PMID: 15365169. Pubmed Central PMCID: PMC518764.
86 Gilbert R, Nalbanoglu J, Tinsley JM, Massie B, Davies KE, Karpati G. Efficient utrophin expression following adenovirus gene transfer in dystrophic muscle. Biochem Biophys Res Commun. 1998;242(1):244–7. PubMed PMID: 9439643.
87 Cerletti M, Negri T, Cozzi F, Colpo R, Andreetta F, Croci D, et al. Dystrophic phenotype of canine X-linked muscular dystrophy is mitigated by adenovirus-mediated utrophin gene transfer. Gene Ther. 2003;10(9):750–7. PubMed PMID: 12704413.
88 Bashir R, Britton S, Strachan T, Keers S, Vafiadaki E, Lako M, et al. A gene related to Caenorhabditis elegans spermatogenesis factor fer-1 is mutated in limb-girdle muscular dystrophy type 2B. Nat Genet. 1998;20(1):37–42. PubMed PMID: 9731527.
89 Liu J, Aoki M, Illa I, Wu C, Fardeau M, Angelini C, et al. Dysferlin, a novel skeletal muscle gene, is mutated in Miyoshi myopathy and limb girdle muscular dystrophy. Nat Genet. 1998;20(1):31–6. PubMed PMID: 9731526.
90 Bansal D, Miyake K, Vogel SS, Groh S, Chen CC, Williamson R, et al. Defective membrane repair in dysferlin-deficient muscular dystrophy. Nature. 2003;423(6936):168–72. PubMed PMID: 12736685.
91 Han R, Campbell KP. Dysferlin and muscle membrane repair. Curr Opin Cell Biol. 2007;19(4):409–16. PubMed PMID: 17662592. Pubmed Central PMCID: PMC2144911.
92 Weiler T, Bashir R, Anderson LV, Davison K, Moss JA, Britton S, et al. Identical mutation in patients with limb girdle muscular dystrophy type 2B or Miyoshi myopathy suggests a role for modifier gene(s). Hum Mol Genet. 1999;8(5):871–7. PubMed PMID: 10196377.
93 Illarioshkin SN, Ivanova-Smolenskaya IA, Greenberg CR, Nylen E, Sukhorukov VS, Poleshchuk VV, et al. Identical dysferlin mutation in limb-girdle muscular dystrophy type 2B and distal myopathy. Neurology. 2000;55(12):1931–3. PubMed PMID: 11134403.
94 Wenzel K, Geier C, Qadri F, Hubner N, Schulz H, Erdmann B, et al. Dysfunction of dysferlin-deficient hearts. J Mol Med (Berl). 2007;85(11):1203–14. PubMed PMID: 17828519.
95 Sinnreich M, Therrien C, Karpati G. Lariat branch point mutation in the dysferlin gene with mild limb-girdle muscular dystrophy. Neurology. 2006;66(7):1114–6. PubMed PMID: 16606933.
96 Aartsma-Rus A, Singh KH, Fokkema IF, Ginjaar IB, van Ommen GJ, den Dunnen JT, et al. Therapeutic exon skipping for dysferlinopathies? Eur J Hum Genet. 2010;18(8):889–94. PubMed PMID: 20145676. Pubmed Central PMCID: PMC2987387.
97 Wang B, Yang Z, Brisson BK, Feng H, Zhang Z, Welch EM, et al. Membrane blebbing as an assessment of functional rescue of dysferlin-deficient human myotubes via nonsense suppression. J Appl Physiol. 2010;109(3):901–5. PubMed PMID: 20558759.
98 Krahn M, Wein N, Bartoli M, Lostal W, Courrier S, Bourg-Alibert N, et al. A naturally occurring human minidysferlin protein repairs sarcolemmal lesions in a mouse model of dysferlinopathy. Sci Transl Med. 2010;2(50):50ra69. PubMed PMID: 20861509.
99 Azakir BA, Di Fulvio S, Salomon S, Brockhoff M, Therrien C, Sinnreich M. Modular dispensability of dysferlin C2 domains reveals rational design for mini-dysferlin molecules. J Biol Chem. 2012;287(33):27629–36. PubMed PMID: 22736764. Pubmed Central PMCID: PMC3431656.
100 Azakir BA, Di Fulvio S, Kinter J, Sinnreich M. Proteasomal inhibition restores biological function of mis-sense mutated dysferlin in patient-derived muscle cells. J Biol Chem. 2012;287(13):10344–54. PubMed PMID: 22318734. Pubmed Central PMCID: PMC3323038.
101 Schoewel V, Marg A, Kunz S, Overkamp T, Carrazedo RS, Zacharias U, et al. Dysferlin-peptides reallocate mutated dysferlin thereby restoring function. PLoS One. 2012;7(11):e49603. PubMed PMID: 23185377. Pubmed Central PMCID: PMC3502493.
102 Brook JD, McCurrach ME, Harley HG, Buckler AJ, Church D, Aburatani H, et al. Molecular basis of myotonic dystrophy: expansion of a trinucleotide (CTG) repeat at the 3' end of a transcript encoding a protein kinase family member. Cell. 1992;68(4):799–808. PubMed PMID: 1310900.
103 Liquori CL, Ricker K, Moseley ML, Jacobsen JF, Kress W, Naylor SL, et al. Myotonic dystrophy type 2 caused by a CCTG expansion in intron 1 of ZNF9. Science. 2001;293(5531):864–7. PubMed PMID: 11486088.
104 de Die-Smulders CE, Howeler CJ, Thijs C, Mirandolle JF, Anten HB, Smeets HJ, et al. Age and causes of death in adult-onset myotonic dystrophy. Brain. 1998;121(Pt 8):1557–63. PubMed PMID: 9712016.
105 Harper PS. Congenital myotonic dystrophy in Britain. I. Clinical aspects. Arch Dis Child. 1975;50(7):505-13. PubMed PMID: 1101835.
106 Reardon W, Harley HG, Brook JD, Rundle SA, Crow S, Harper PS, et al. Minimal expression of myotonic dystrophy: a clinical and molecular analysis. J Med Genet. 1992;29(11):770–3. PubMed PMID: 1453424.
107 MacDonald JR, Hill JD, Tarnopolsky MA. Modafinil reduces excessive somnolence and enhances mood in patients with myotonic dystrophy. Neurology. 2002;59(12):1876–80. PubMed PMID: 12499477.
108 Damian MS, Gerlach A, Schmidt F, Lehmann E, Reichmann H. Modafinil for excessive daytime sleepiness in myotonic dystrophy. Neurology. 2001;56(6):794–6. PubMed PMID: 11274321.
109 Brunner HG, Bruggenwirth HT, Nillesen W, Jansen G, Hamel BC, Hoppe RL, et al. Influence of sex of the transmitting parent as well as of parental allele size on the CTG expansion in myotonic dystrophy (DM). Am J Hum Genet. 1993;53(5):1016–23. PubMed PMID: 8213829.
110 Jansen G, Willems P, Coerwinkel M, Nillesen W, Smeets H, Vits L, et al. Gonosomal mosaicism in myotonic dystrophy patients: involvement of mitotic events in (CTG)n repeat variation and selection against extreme expansion in sperm. Am J Hum Genet. 1994;54(4):575–85. PubMed PMID: 8128954.
111 Kovtun IV, McMurray CT. Trinucleotide expansion in haploid germ cells by gap repair. Nat Genet. 2001;27(4):407–11. PubMed PMID: 11279522.
112 Fu YH, Friedman DL, Richards S, Pearlman JA, Gibbs RA, Pizzuti A, et al. Decreased expression of myotonin-protein kinase messenger RNA and protein in adult form of myotonic dystrophy. Science. 1993;260(5105):235–8. PubMed PMID: 8469976.
113 Reddy S, Smith DB, Rich MM, Leferovich JM, Reilly P, Davis BM, et al. Mice lacking the myotonic dystrophy protein kinase develop a late onset progressive myopathy. Nat Genet. 1996;13(3):325–35. PubMed PMID: 8673132.
114 Jansen G, Groenen PJ, Bachner D, Jap PH, Coerwinkel M, Oerlemans F, et al. Abnormal myotonic dystrophy protein kinase levels produce only mild myopathy in mice. Nat Genet. 1996;13(3):316–24. PubMed PMID: 8673131.
115 Mankodi A, Logigian E, Callahan L, McClain C, White R, Henderson D, et al. Myotonic dystrophy in transgenic mice expressing an expanded CUG repeat. Science. 2000;289(5485):1769–73. PubMed PMID: 10976074.
116 Michalowski S, Miller JW, Urbinati CR, Paliouras M, Swanson MS, Griffith J. Visualization of double-stranded RNAs from the myotonic dystrophy protein kinase gene and interactions with CUG-binding protein. Nucleic Acids Res. 1999;27(17):3534–42. PubMed PMID: 10446244.
117 Mooers BH, Logue JS, Berglund JA. The structural basis of myotonic dystrophy from the crystal structure of CUG repeats. Proc Natl Acad Sci U S A. 2005;102(46):16626–31. PubMed PMID: 16269545.
118 Kino Y, Mori D, Oma Y, Takeshita Y, Sasagawa N, Ishiura S. Muscleblind protein, MBNL1/EXP, binds specifically to CHHG repeats. Hum Mol Genet. 2004;13(5):495–507. PubMed PMID: 14722159.
119 Miller JW, Urbinati CR, Teng-Umnuay P, Stenberg MG, Byrne BJ, Thornton CA, et al. Recruitment of human muscleblind proteins to (CUG)(n) expansions associated with myotonic dystrophy. Embo J. 2000;19(17):4439–48. PubMed PMID: 10970838.
120 Faustino NA, Cooper TA. Pre-mRNA splicing and human disease. Genes Dev. 2003;17(4):419–37. PubMed PMID: 12600935.
121 Kanadia RN, Johnstone KA, Mankodi A, Lungu C, Thornton CA, Esson D, et al. A muscleblind knockout model for myotonic dystrophy. Science. 2003;302(5652):1978–80. PubMed PMID: 14671308.
122 Kuyumcu-Martinez NM, Wang GS, Cooper TA. Increased steady-state levels of CUGBP1 in myotonic dystrophy 1 are due to PKC-mediated hyperphosphorylation. Mol Cell. 2007;28(1):68–78. PubMed PMID: 17936705. Pubmed Central PMCID: PMC2083558.
123 Ho TH, Bundman D, Armstrong DL, Cooper TA. Transgenic mice expressing CUG-BP1 reproduce splicing mis-regulation observed in myotonic dystrophy. Hum Mol Genet. 2005;14(11):1539–47. PubMed PMID: 15843400.
124 Mulders S, van den Broek W, Wheeler T, Croes H, van Kuik-Romeijn P, de Kimpe S, et al. Triplet-repeat oligonucleotide-mediated reversal of RNA toxicity in myotonic dystrophy. Proc Natl Acad Sci U S A. 2009;106(33):13915-20. PubMed PMID: 19667189. Pubmed Central PMCID: PMC2728995.
125 Wheeler T, Sobczak K, Lueck J, Osborne R, Lin X, Dirksen R, et al. Reversal of RNA dominance by displacement of protein sequestered on triplet repeat RNA. Science. 2009;325(5938):336–9. PubMed PMID: 19608921.
126 Lee JE, Bennett CF, Cooper TA. RNase H-mediated degradation of toxic RNA in myotonic dystrophy type 1. Proc Natl Acad Sci U S A. 2012;109(11):4221–6. PubMed PMID: 22371589. Pubmed Central PMCID: PMC3306674.
127 Wheeler TM, Leger AJ, Pandey SK, MacLeod AR, Nakamori M, Cheng SH, et al. Targeting nuclear RNA for in vivo correction of myotonic dystrophy. Nature. 2012;488(7409):111–5. PubMed PMID: 22859208.
128 Furling D, Doucet G, Langlois MA, Timchenko L, Belanger E, Cossette L, et al. Viral vector producing antisense RNA restores myotonic dystrophy myoblast functions. Gene Ther. 2003;10(9):795–802. PubMed PMID: 12704419.
129 Langlois MA, Boniface C, Wang G, Alluin J, Salvaterra PM, Puymirat J, et al. Cytoplasmic and nuclear retained DMPK mRNAs are targets for RNA interference in myotonic dystrophy cells. J Biol Chem. 2005;280(17):16949–54. PubMed PMID: 15722335.
130 Langlois MA, Lee NS, Rossi JJ, Puymirat J. Hammerhead ribozyme-mediated destruction of nuclear foci in myotonic dystrophy myoblasts. Mol Ther. 2003;7(5 Pt 1):670-80. PubMed PMID: 12718910.
131 García-López A, Llamusí B, Orzáez M, Pérez-Payá E, Artero RD. In vivo discovery of a peptide that prevents CUG-RNA hairpin formation and reverses RNA toxicity in myotonic dystrophy models. Proc Natl Acad Sci U S A. 2011;108(29):11866–71. PubMed PMID: 21730182. Pubmed Central PMCID: PMC3141925.
132 Pushechnikov A, Lee M, Childs-Disney J, Sobczak K, French J, Thornton C, et al. Rational design of ligands targeting triplet repeating transcripts that cause RNA dominant disease: application to myotonic muscular dystrophy type 1 and spinocerebellar ataxia type 3. J Am Chem Soc. 2009;131(28):9767–79. PubMed PMID: 19552411. Pubmed Central PMCID: PMC2731475.
133 Childs-Disney JL, Hoskins J, Rzuczek SG, Thornton CA, Disney MD. Rationally designed small molecules targeting the RNA that causes myotonic dystrophy type 1 are potently bioactive. ACS Chem Biol. 2012;7(5):856–62. PubMed PMID: 22332923. Pubmed Central PMCID: PMC3356481.
134 Warf M, Nakamori M, Matthys C, Thornton C, Berglund J. Pentamidine reverses the splicing defects associated with myotonic dystrophy. Proc Natl Acad Sci U S A. 2009;106(44):18551–6. PubMed PMID: 19822739. Pubmed Central PMCID: PMC2774031.
135 Lunt PW, Harper PS. Genetic counselling in facioscapulohumeral muscular dystrophy. J Med Genet. 1991;28(10):655–64. PubMed PMID: 1941962.
136 Jardine PE, Koch MC, Lunt PW, Maynard J, Bathke KD, Harper PS, et al. De novo facioscapulohumeral muscular dystrophy defined by DNA probe p13E-11 (D4F104S1). Arch Dis Child. 1994;71(3):221–7. PubMed PMID: 7979495.
137 Wohlgemuth M, van der Kooi EL, van Kesteren RG, van der Maarel SM, Padberg GW. Ventilatory support in facioscapulohumeral muscular dystrophy. Neurology. 2004;63(1):176–8. PubMed PMID: 15249635.
138 Wohlgemuth M, de Swart BJ, Kalf JG, Joosten FB, Van der Vliet AM, Padberg GW. Dysphagia in facioscapulohumeral muscular dystrophy. Neurology. 2006;66(12):1926–8. PubMed PMID: 16801662.
139 Fitzsimons RB, Gurwin EB, Bird AC. Retinal vascular abnormalities in facioscapulohumeral muscular dystrophy. A general association with genetic and therapeutic implications. Brain. 1987;110 ( Pt 3):631–48. PubMed PMID: 3580827.
140 Padberg GW, Frants RR, Brouwer OF, Wijmenga C, Bakker E, Sandkuijl LA. Facioscapulohumeral muscular dystrophy in the Dutch population. Muscle Nerve. 1995;2:S81–4. PubMed PMID: 7739631.
141 Wijmenga C, Padberg GW, Moerer P, Wiegant J, Liem L, Brouwer OF, et al. Mapping of facioscapulohumeral muscular dystrophy gene to chromosome 4q35-qter by multipoint linkage analysis and in situ hybridization. Genomics. 1991;9(4):570–5. PubMed PMID: 2037288.
142 van Deutekom JC, Wijmenga C, van Tienhoven EA, Gruter AM, Hewitt JE, Padberg GW, et al. FSHD associated DNA rearrangements are due to deletions of integral copies of a 3.2 kb tandemly repeated unit. Hum Mol Genet. 1993;2(12):2037–42. PubMed PMID: 8111371.
143 van Geel M, Dickson MC, Beck AF, Bolland DJ, Frants RR, van der Maarel SM, et al. Genomic analysis of human chromosome 10q and 4q telomeres suggests a common origin. Genomics. 2002;79(2):210–7. PubMed PMID: 11829491.
144 Lemmers RJ, de Kievit P, Sandkuijl L, Padberg GW, van Ommen GJ, Frants RR, et al. Facioscapulohumeral muscular dystrophy is uniquely associated with one of the two variants of the 4q subtelomere. Nat Genet. 2002;32(2):235–6. PubMed PMID: 12355084.
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