Animal Models of Muscular Dystrophy

Abstract

The muscular dystrophies (MDs) represent a diverse collection of inherited human disorders, which affect to varying degrees skeletal, cardiac, and sometimes smooth muscle (Emery, 2002¹). To date, more than 50 different genes have been implicated as causing one or more types of MD (Bansal et al., 2003²). In many cases, invaluable insights into disease mechanisms, structure and function of gene products, and approaches for therapeutic interventions have benefited from the study of animal models of the different MDs (Arnett et al., 2009³). The large number of genes that are associated with MD and the tremendous number of animal models that have been developed preclude a complete discussion of each in the context of this review. However, we summarize here a number of the more commonly used models together with a mixture of different types of gene and MD, which serves to give a general overview of the value of animal models of MD for research and therapeutic development.

I. Dystrophin

Duchenne muscular dystrophy (DMD) and the milder Becker MD (BMD) are caused by dystrophin gene mutations.^4,5 DMD/BMD is thought to be the most common form of muscular dystrophy (MD), and the most common lethal genetic disorder of children.⁶ Full-length dystrophin is expressed in all muscles, while shorter, truncated isoforms are expressed in many other tissues.⁷ Dystrophin is part of a large complex of integral and peripheral proteins that link the γ-actin cytoskeleton to the extracellular matrix via the dystrophin–glycoprotein complex (DGC) (Fig. 1).⁸ Studies of dystrophin and DMD/BMD have been greatly facilitated by the availability of numerous murine and canine models of DMD.⁹

Fig. 1.

The dystrophin–glycoprotein complex (DGC) contains many proteins that, when not expressed, lead to various forms of muscular dystrophy. Shown is the organization of the DGC in skeletal muscle. Dystrophin is defective in DMD and BMD; α-, β-, γ-, and δ- sarcoglycans (Sgn) are defective in four different types of LGMD; laminin α2 is defective in one form of congenital muscular dystrophy; collagen VI in the extracellular matrix is defective in Bethlem myopathy and Ullrich congenital muscular dystrophy; while numerous other cytosolic- and membrane-associated proteins lead to other MDs. Other abbreviations include Syn, syntrophin; SPN, sarcospan; Dgn, dystroglycan; nNOS, neuronal nitric oxide synthase. Dystrophin domains listed include the N-terminal actin-binding domain (NT), various hinges (H), a WW domain (W), a cysteine-rich domain (CR), and a C-terminal domain (CT). Orange and red balls in dystrophin represent spectrin-like repeats, orange ones are acidic, while blue ones are basic and contribute to actin binding.

A. Mouse Models for Duchenne Muscular Dystrophy

Mouse models have been used extensively to elucidate the pathogenic mechanism of DMD and have been indispensable in the development of therapeutic approaches. Many mouse models with mutations in the DMD gene have been identified. Among these models, the mdx mouse is the most commonly used to analyze dystrophin gene expression and function. The mdx mouse contains a premature stop codon in exon 23 that leads to loss of full-length dystrophin, although smaller isoforms are still expressed.^10–12 Although mdx skeletal muscle shows moderate signs of dystrophy, young mice exhibit little weakness and live 80% as long as controls, significantly more than that of DMD patients.¹³ This mild, yet progressive phenotype has been attributed in part to compensatory upregulation of utrophin.^14,15 Histological examination of mdx muscle during various stages of development reveals that muscle fiber necrosis and cellular infiltration begin at approximately 3 weeks of age. This is followed by a crises period that peaks at approximately 4–6 weeks of age and is characterized by the presence of extensive necrosis, regenerating muscle fibers with centrally located nuclei, and elevated levels of serum creatine kinase (CK).^10,16 After 12 weeks, the cycles of necrosis and regeneration begin to slow, although necrotic myofibers are present for the remainder of their lifespan. The fibrosis and infiltration of inflammatory cells in skeletal and cardiac muscle of the mdx are much milder than that observed in DMD patients.^17,18 In contrast, the mdx mouse diaphragm exhibits severe pathological changes and functional deficit comparable to that of DMD limb muscle.^19,20

Four additional strains of mdx mice, mdx^2cv–5cv, have been generated with N-ethylnitrosourea chemical mutagenesis.²¹ All these strains have point mutations that lead to loss of full-length dystrophin isoforms. The relative location of these mutations results in a series of mdx mouse mutants that vary in their expression of different dystrophin isoforms, such that these mice should be useful for analysis of the role of the various isoforms.¹² Despite their differences, all five mdx strains display essentially identical muscle pathology as mdx mice, although additional phenotypes have been observed.^21–23 In the mdx^2cv mouse, a single base change in the splice acceptor site of dystrophin intron 42 generates multiple transcripts due to aberrant splicing that eliminates the normal open reading frame. The mdx^3cv allele arises from a mutant splice acceptor site in intron 65,^12,21,22 leading to severe loss of most dystrophin isoforms, including Dp71. Of the five original mdx strains, only the mdx^3cv mice accumulate detectable levels of dystrophin-positive muscle fibers, although the expressed dystrophin lacks a key functional domain and is largely nonfunctional.^22–24 The mutation in the mdx^4cv mouse is a point mutation that creates a stop codon in exon 53, whereas in the mdx^5cv allele, a transversion mutation in exon 10 creates a new splice donor that generates a frame-shifting deletion in the mRNA. Both the mdx^4cv and mdx^5cv strains display a low background of reverent dystrophin-containing fibers, making them particularly useful strains in gene transfer studies exploring the feasibility of DMD therapy.^23,25,26

In the mdx^βgeo mouse, a β-galactosidase/neomycin phosphotransferase (βgeo) reporter gene is inserted 3′ to exon 63 of the dystrophin gene, affecting translation of dystrophin isoforms, including Dp71.²⁷ These mice develop a dilated esophagus, cardiac hypertrophy, and the same skeletal muscle pathology as mdx mice. The lacZ reporter gene of mdx^βgeo allows detection of dystrophin expression by staining for β-galactosidase activity.

Another mutant mouse known as mdx52 was generated by a homologous recombination technique.²⁸ In this model, exon 52 of the dystrophin gene is disrupted, and dystrophin along with shorter dystrophin isoforms (Dp140 and Dp260) are absent. This exon was targeted to replicate the most common out-of-frame deletion mutation that occurs in 13% of DMD patients.^29–31 Recently, antisense-mediated skipping of exon 51 using antisense oligonucleotides (AOs) to restore the reading frame of the dystrophin gene was found to be successful in some patients with DMD.^32–34 Exon 51 skipping using AO in mdx52 mice successfully converted an out-of-frame mutation into an in-frame mutation, resulting in amelioration of the dystrophic pathology and improved muscle function.³⁵

Hanaoka and colleagues generated a new DMD mouse model known as DMD-null. In this model, the entire genomic region of the DMD gene on mouse chromosome X was deleted using a Cre-loxP recombination system.³⁶ These DMD-null males exhibited some behavioral abnormality, sterility, and pathological features very similar to those of the mdx mice. As the production of all dystrophin isoforms is prevented in the DMD-null mouse, this model system may be used to facilitate studies to clarify the function of individual dystrophin isoforms.

In efforts to make the mdx muscle phenotype more similar to that of patients, several additional mutations have been crossed onto the mdx background to generate double knockouts (DKOs). The most widely used is a dystrophin:utrophin DKO (mdx:utrn^−/−).^37,38, These mice display a severe phenotype and live on average only about 3 months. The severity of the phenotype supports the concept that utrophin upregulation in dystrophic muscles partially compensates for the absence of dystrophin. Further, the mdx:utrn^−/− have proved useful in gene therapy studies, where the phenotype can be largely eliminated by muscle-specific expression of utrophin, miniutrophin, or mini- or micro-dystrophin.^{25,26,37–40} Two newer DKO strains have recently been generated that not only resulted in a several phonotype but also rely on mutations that appear to faithfully reproduce the human condition. The first is a cross between mdx and a mutation of cytidine monophosphate–sialic acid hydroxylase (Cmah).⁴¹ Cmah is inactive in humans due to a mutation, and it prevents glycosylation with N-glycolylneuraminic acid. The second is an mdx:mTR cross that effectively lacks telomerase activity.⁴² This cross becomes severe due to the impaired ability of myogenic stem cells to support ongoing muscle regeneration.

B. Canine Models for DMD

Clinical diagnosis of MD has been observed in the dog for several decades. Identifying a particular MD with broad symptoms (i.e., weakness and intolerance to exercise) has proven difficult. With the increasing availability of research tools, more human MDs have found a pathologically similar model in canines. Yet, the infrequency of these diagnoses and the difficulties in identifying and propagating the mutations by breeding have hindered multiple MD strains from being used as valuable research tools. One MD that has been successfully generated and utilized throughout the world is the X-linked dystrophin-deficient dog. Spontaneous mutations causing dystrophinopathy have been identified in several breeds of dog.^43–45 Multiple colonies of the golden retriever muscular dystrophy dog (GRMD) have been created and is the most extensively studied breed for this model, but it is not the only one.^9,46 Due to prior use in research and its smaller size, the GRMD mutation has been bred onto the beagle background.^47,48 Severe symptoms commonly appear at 6 months of age in the GRMD dog, but unlike the mdx mouse, the degree of severity and time of progression are quite variable. Pathology has been observed in utero and massive necrosis can result in neonatal death.⁴⁶ Compounded by the increased cost of maintaining a larger species, the GRMD model can be problematic. However, use of the GRMD model has gained much emphasis with its more clinically similar pathology than the mdx mouse model.

II. The Sarcoglycans

As a clinical group, the MDs were first appreciated in the mid-1800s with a detailed description of DMD.^49,50 Approximately a century following, another group of clinically comparable progressive MDs distinct from DMD/BMD muscular dystrophy emerged which were found to follow an autosomal recessive mode of inheritance. Given their phenotypic similarity, these diseases were originally termed Duchenne-like autosomal recessive MD or severe childhood autosomal recessive MD.⁵¹ Several decades later, the discovery of mutations in the dystrophin gene as the genetic cause of DMD,⁵² subsequent description of the DGC,⁵³ and demarcation of the striated muscle transmembrane sarcoglycan complex paved the way for initial characterization of the sarcoglycanopathies. Four genetically related MDs arose with pathological mutations in genes for alpha-sarcoglycan (α-SG),⁵⁴ beta-sarcoglycan (β-SG),⁵⁵ gamma-sarcoglycan (γ-SG),⁵⁶ and delta-sarcoglycan (δ-SG)⁵⁷ defining these distinct autosomal recessive MDs. Given that these disorders primarily affected proximal muscles around the scapular and pelvic girdles, and elevation of the genetic classification of limb-girdle muscular dystrophies (LGMD), there was the assignment of LGMD2C-F for the genes representing γ-SG, α-SG, β-SG, and δ-SG, respectively.

A. δ-Sarcoglycan and LGMD2F

The BIO14.6 hamster strain was established in 1962 and is recognized as the first animal model of sarcoglycanopathies.⁵⁸ However, it was not until after the discovery of the causative mutation in LGMD2F patients⁵⁹ that a large deletion in the 5′ portion of the δ-SG gene was identified.⁶⁰ The hamsters die prematurely (∼ 8 months) from progressive myocardial necrosis and ultimately heart failure. Compared with skeletal muscle, damage to cardiac muscle occurs early (∼ 5 weeks), consistent with a cardiomyopathic phenotype with decreased stroke volume, cardiac output, and increased ventricular hypertrophy.⁶¹ As the original BIO14.6 strain was identified, there have been at least three additional strains developed: the J2-NK, the CHF147 (formerly VMX7.1), and the TO-2 strain. Differences, such as the expression profile of individual sarcoglycan isoforms, have been observed in the TO-2 as compared to the BIO14.6 model. For example, in the BIO14.6 muscles, residual protein levels of α-SG and γ-SG can be detected on the sarcolemma, while in the TO-2, there is a complete absence of these isoforms. Further, the TO-2 has a shorter lifespan with a tendency to develop a more severe cardiomyopathy displaying a necrotic mode of pathology, while the BIO14.6 model displays a hypertrophic cardiomyopathy.^62,63 Mice deficient in δ-SG have also been generated with subsequent complete loss of the entire SG complex,^64,65a and as with the BIO14.6 hamster, these mice display classical histological findings of dystrophy inclusive of muscle degeneration, necrosis, inflammation, fibrosis, and reduced survival. However, muscle mass, peak twitch tension (P₁) and peak tetanic tension (P₀), and specific force (P₀/CSA) do not differ from wild-type muscle with the latter being in contrast to the BIO14.6 hamster^65b suggesting species-specific variations in pathophysiology. Similar to the BIO14.6 hamster model, these mice develop cardiomyopathy with a relatively early onset (∼ 8 weeks) with areas of fibrosis. By 32 weeks, a detrimental change in pathology is observed with concurrent reduction in ejection fraction, and evidence of right ventricular dilation suggestive of pulmonary dysfunction.^65c Thus, similar to the hamster model, the δ-SG-null mouse reproduces many of the pathophysiological features of the human disease, providing a useful preclinical research model.

B. γ-Sarcoglycan and LGMD2C

Mice with a targeted deletion of γ-SG also exhibit progressive MD and cardiomyopathy with a subsequent destabilization of β-SG and δ-SG while α-SG is highly reduced.⁶⁶ Like the δ-SG-null hamster, cardiomyopathy in γ-SG-null mice may be dilated or hypertrophic.^62,66 Similar to findings in LGMD2C patients, membrane disruptions are a common feature of mice deficient for γ-SG as evidenced by Evans blue dye uptake and elevation of the muscle-specific enzyme M-CK in serum.^66,67 Importantly, in striated muscle, the expression of dystrophin, dystroglycan, and laminin appears unaltered with the loss of γ-SG when evaluated by immunostaining.⁶⁴ Thus, the mechanical linkage between the extracellular matrix and the intracellular cytoskeleton appears to be maintained in the absence of γ-SG. Further, γ-SG-deficient mice show normal resistance to mechanical strain and no evidence of contraction-induced injury after exercise when compared to its wild-type counterparts.⁶⁶ These results are similar to the BIO14.6 hamster and in contrast to muscles of α-SG, β-SG, and δ-SG-null mice (see below). Considering the sarcoglycans are secondarily reduced in dystrophin-deficient mice, this advocates that nonmechanical properties contribute to the pathophysiology of MD in genetically varied forms of disease.

C. β-Sarcoglycan and LGMD2E

Mice deficient for β-SG have also been generated which, like δ-SG-null mice, display MD with large areas of necrosis/fibrosis and cardiomyopathy.^68,69 As an indicator of acute myocardial necrosis preceding that of fibrotic lesion formation, serum levels of the cardiac-specific troponin-I have been found elevated in these mice.^65a,68,70 Additionally, like δ-SG-null mice, β-SG-null mice show severely reduced expression of the entire sarcoglycan complex inclusive of sarcospan, increased muscle mass, decreased specific force generation, myofiber cross-sectional area, and resistance to contraction-induced injury.^68,69,71 In support of the former, coimmunoprecipitation studies have shown a strong association between β-SG and δ-SG, proposed to form a core complex enabling delivery and retention of the sarcoglycan complex to the sarcolemma.⁷² Correlative support of this hypothesis can be found in human patients with either β-SG or δ-SG mutations, where there is a strong reduction or complete absence of the entire sarcoglycan complex.^73–75

D. α-Sarcoglycan and LGMD2D

Mice lacking α-SG have been generated by targeted disruption of the α-SG gene.⁷⁶ These mice show a complete loss of the sarcoglycan complex, demonstrate a progressive MD, and like LGMD2D patients present with an absence of cardiomyopathy.⁷⁶ Like the γ-, β-, and δ-SG-null mice, the α-SG-null mice demonstrate membrane permeability defects as evidenced by Evans blue dye uptake and elevated serum CK. Given this similarity across dystrophic models, it remains unclear whether or not changes in increased membrane permeability are causative for muscle degeneration or rather a result of it. A further interesting aspect of α-SG-null mice that differs from the γ-SG-null mice is an apparent decrease in expression of α-dystroglycan (αDG) and dystrophin with an inherent increase in expression of utrophin at the sarcolemma.⁷⁶ The molecular basis for these differences has not been established, though it is tempting to speculate that the residual amount of α-SG present in the γ-SG-null could be sufficient to allow normal levels of dystrophin and dystroglycan. In the α-SG-null mice, there are notable muscle-specific differences in contractile properties. For example, in hypertrophic slow-twitch soleus muscle, there is a demonstrated increase in maximal force values and relatively unchanged specific force relative to wild-type controls.⁷⁶ However, in fast-twitch extensor digitorum longus (EDL) muscles, maximal force values are unchanged while specific force is significantly lower than wild-type controls.⁷⁶ In diaphragm of α-SG-null mice, there is a significant reduction in maximal force, coupling between longitudinal and transverse properties, and muscle extensibility in the direction of muscle fibers and transverse to fibers.⁷⁷ Thus, as a whole, the sarcoglycan complex may also provide a complex mechanical function in the diaphragm by influencing the properties of muscle stiffness, muscle viscoelasticity, and modulation of the contractile properties.

E. Canine Models of Sarcoglycan Deficiency

Three breeds of dogs have also been reported with deficiencies in expression of the sarcoglycan complex in skeletal muscle.^78,79 Where the mutation lies in each of these cases has yet to be determined, but as discussed above for both rodents and humans, a deficiency in one sarcoglycan can destabilize the subcomplex made by all four of these isoforms.

III. Calcium and Dystrophic Pathology

A. Calcium Contributes to the Pathology of Dystrophic Skeletal Muscles

Calcium plays a dual role in muscle function, serving as a signaling molecule as well as a regulator of muscle contraction and relaxation.⁸⁰ To effectively regulate cytosolic [Ca²⁺], muscle fibers contain a complex system of channels, pumps, and buffering proteins, each dedicated to mobilize, transport, or sequester Ca²⁺.⁸⁰ In dystrophin-deficient muscle, all three components, that is, channels, pumps, and buffering proteins, appear to be dysfunctional. Ca²⁺ channels in the sarcolemma of mdx mice exhibit increased activity,^81,82 while Ca²⁺ buffering proteins^83,84 and ATPase pumps in the sarcoplasmic reticulum^85,86 demonstrate an impaired ability to sequester Ca²⁺. Together, the compromised Ca²⁺-cycling system in mdx muscles results in an elevated level of resting intracellular [Ca²⁺].^87–89 Accumulation of Ca²⁺ is particularly deleterious to the survivability of muscle fibers, as Ca²⁺ activates a variety of free radical and enzymatic processes that target the cytoskeleton, sarcolemma, and mitochondria. Left unchecked, these processes may ultimately result in the necrosis of the muscle fiber.⁹⁰

Investigators have gained valuable insight into the relationship between calcium and MD by developing several mouse models with altered calcium handling properties. One class of calcium handling proteins to be overexpressed is the transient receptor potential canonical (TRP) channels that facilitate the entry of Ca²⁺ into muscle fibers. Overexpression of TRP channels in wild-type mice causes skeletal muscles to exhibit increased fibrosis, immune cell infiltration, fatty tissue, and continuous cycles of muscle regeneration. This mouse model possesses many of the pathological characteristics of dystrophic mice and demonstrates that calcium misregulation alone is sufficient to trigger a MD-like phenotype. In a related set of experiments, investigators were able to partially correct the pathology of dystrophic mdx mice by overexpressing nonfunctional mutant TRP channels.^91,92 The rescue effect stems from the suppression of endogenous TRP channel activity elicited by the dominant-negative mutant TRP channels.

B. Dysferlin Gene Mutations

In addition to overactive calcium channels, compromised membrane-repair processes are also capable of eliciting MD. Dysferlin is a protein that augments membrane repair^2,93,94 and deficiencies have been linked to LGMD2B and Miyoshi myopathy.^95,96 In mouse models that lack dysferlin, skeletal muscles suffer from a progressive dystrophy and display altered morphology, muscle necrosis, macrophage infiltration, and fatty tissue replacement.^2,97 Without dysferlin, lesions in the membrane that may occur during contractile activity remain unrepaired, allowing calcium to rush into the muscle fiber. The uncontrolled influx of calcium into the cytosol is likely to overwhelm the endogenous calcium handling apparatus and trigger gradual muscle fiber necrosis. Studies on these mouse models suggest that imbalances in cytosolic calcium levels are sufficient to induce progressive MD, independent of the absence or presence of structural proteins such as dystrophin and the DGC. Other recent studies also suggest that dysferlin may be a component of T-tubules, but whether it regulates calcium flux there is not clear.^93,98

C. Calpain 3 and LGMD2A

Mutations in calpain 3 gene cause LGMD2A, one of the most prevalent forms of the disease.^99–101 Calpain 3 is a calcium-dependent nonlysosomal protease involved primarily in regulating sarcomere formation and remodeling.¹⁰² Additionally, calpain 3 serves structural functions as a component of the skeletal muscle triad¹⁰³ and the dysferlin complex.¹⁰⁴ Although calpain 3 has been found in many subcellular locations, the majority of it is anchored to titin, a giant sarcomeric protein that acts as a molecular ruler and a template for sarcomere assembly during muscle cell development.^105–107 As mutations in titin result in LGMD2J and a secondary deficiency of calpain 3,¹⁰⁸ mouse models for both forms of the disease will be discussed together.

Two calpain 3 knockout mouse models have been created. Richard et al. generated mice with disrupted proteolytic site of calpain 3 by substituting exons 2 and 3 with a neoR cassette using homologous recombination.¹⁰⁹ Skeletal muscles from the transgenic mice showed 80% decrease in calpain 3 mRNA and a complete absence of calpain 3 protein. Phenotypically, the muscles revealed regions of focal necrosis and apoptosis of nuclei associated with perturbation of NF-kB/IkBα as well as disruption of sarcolemmal integrity. Spencer's group took a different approach and generated calpain 3-deficient mice by utilizing a gene trap retroviral vector that introduced premature stop signals that completely abolished mRNA and protein expression.¹¹⁰ The phenotype more closely resembled LGMD2A^111,112 as it showed muscle necrosis in combination with fiber atrophy. The studies utilizing these mice indicated that calpain 3 participates in remodeling of the sarcomere by promoting ubiquitination of its ligands.¹⁰² Further, calpain 3-deficient mice were shown to have reduced calcium release in fibers due to perturbations in the triad-associated complex.¹⁰³

Although most cases of LGMD2A result from mutations in the proteolytic region of calpain 3, pathogenic mutations are present along the entire sequence of calpain 3 gene. Mice expressing a proteolytically inactive mutant form (C129S) that retains its conformational properties enabled dissecting enzymatic and structural functions of calpain 3. Tagawa et al. generated transgenic mice expressing the mutated protein on the wild-type background.¹¹³ Despite low expression levels, the aged mice showed signs of a mild myopathy, including reduced motor functions, as well as the presence of centrally nucleated nuclei, and lobulated and split fibers. In a more recent study,¹¹⁴ analyses of C129S knock-in mice revealed progressive MD associated with inefficient redistribution of the mutant protein in stretched sarcomeres. Nevertheless, the mice showed a less severe phenotype than calpain 3 knockout mice, presumably due to preserved calpain 3 structural functions in the sarcoplasmic reticulum.^103,115 Another study showed that impaired titin–calpain 3 interaction results in reduced concentration of calpain 3 in the myofibrillar fraction along with its rapid degradation.¹¹⁶ Here, generated transgenic mice expressed mutant cal-pains known to result in LGMD2A with preserved proteolytic function but mutated titin-binding site on the caveolin 3 knockout background.

D. Titin and LGMD2J

Mutations in the last 2 coding exons of titin lead to a mild late onset dominant tibial muscular dystrophy and a severe recessive LGMD2J. The knock-in mice carrying the most common mutation (FINmaj) recapitulate the human pathology, with heterozygotes showing a very mild phenotype and homozygotes developing a progressive MD.¹¹⁷ This mouse model reveals also a secondary reduction in calpain 3 expression levels. Importantly, when the mice were bred onto the calpain 3-deficient background, the pathology of the FINmaj knock-in mice was attenuated, indicating that disrupted regulation of calpain 3 is responsible for the pathology of LGMD2J. It is also important to note that the pathology of another mouse model for titinopathy, severely affected MD with myositis (mdm), was not linked to deregulation of calpain 3.¹¹⁸ The Mdm causing deletion of 83 amino acids in the N2A region of titin arose spontaneously in the Jackson laboratory in the early 1980s.¹¹⁹ The mutation disrupts the calpain 3 binding site in the N2A region of titin. Despite that, mdm/calpain 3-deficient mice showed no change in the mdm pathology.

IV. α-Actinin

The α-actinins are a family of actin-binding proteins that are structurally related to dystrophin. Of the four members in the α-actinin family, only the sarcomeric isoforms α-actinin-2 and α-actinin-3 (Actn3) have been shown to bind dystrophin and other Z-disk associated proteins at the sarcomere,¹²⁰ while the cytoskeletal isoforms α-actinin-1 and α-actinin-4 are involved in cytoskeleton organization and cellular attachment to the extracellular matrix at focal adhesions.¹²¹ As α-actinin-2 is the sole sarcomeric α-actinin isoform expressed in the heart, mutations in α-actinin-2 result in hypertrophic cardiomyopathy.¹²² In contrast, complete deficiency of α-actinin-3 occurs in 16% of the normal human population¹²³ and is associated with reduced muscle strength and power in elite athletes^124–127 and nonathletes.^128,129 Phenotypic analysis of the Actn3 knockout mouse showed that absence of α-actinin-3 alters the structural, metabolic, and physiological characteristics of fast-twitch muscle fibers toward that of slow-twitch fibers,¹³⁰ suggesting that α-actinin-3 expression is essential for optimal rapid power generation in muscles of humans and mice. Recently, α-actinin-3 deficiency has also been associated with increased susceptibility to contraction-induced injury.¹³¹ Work is ongoing to examine the influence of ACTN3 genotype on muscle disease phenotypes by crossing the Actn3 knockout mouse with mouse models of muscle disease, and to discover the therapeutic potential of α-actinin-3 overexpression by generation of Actn3 transgenic models.

V. Fukutin-Related Protein and Dystroglycan Processing

Fukutin-related protein (FKRP) is a putative glycosyltransferase and is one of many that glycosylate αDG, allowing it to stabilize the sarcomere at the basal lamina via binding to laminin α2. Mutations in FKRP cause a broad spectrum of clinical phenotypes with variable disease severity. The most severe forms result in structural brain abnormalities (Walker–Warburg syndrome, WWS and muscle–eye–brain disease, MEB)¹³² and an inabiity to walk (MDC1C).¹³³ FKRP mutations also result in LGMD2I, one of the most common forms of LGMD in childhood. LGMD2I itself is variable—one version has an early onset, Duchenne-like phenotype (severe LGMD2I) and the other a milder, slower progression (mild LGMD2I). Clinical severity has been correlated to depletion of glycosylated αDG and secondary reduction in laminin α2 as a consequence of reduced FKRP expression.^134,135

The effect of FKRP downregulation has been explored in zebrafish¹³⁶ and mice¹³⁷ injected with AOs. While mouse models based on patient FKRP mutations that result in MEB (Y307N),¹³⁸ WWS (E310del), and MDC1C (P448L)¹³⁹ have also been generated, only those retaining the neomycin cassette succeeded in reducing Fkrp transcript levels and recapitulating the severity of the disease phenotypes.^138,139 Of the three mouse models, the E310del was embryonically lethal, FKRP-neo-Y307N resulted in perinatal lethality, while some FKRP-neo-P448L mutant mice survived to 10 months. The difference in lifespan of these models could be associated with the level of Fkrp mRNA/protein present. Homozygous E310del mice completely lack functional FKRP due to deletion of the C-terminus of FKRP that contains a consensus D × D motif commonly found in glycosyltransferases, suggesting that the presence of at least some FKRP activity is critical for embryonic development.¹³⁹ In contrast, the FKRP-neo-Y307N homozygotes expressed approximately 40% of Fkrp mRNA compared to controls,¹³⁸ while FKRP-neo-P448L homozygotes expressed approximately 55%.¹³⁹ This is consistent with results from knockdown of Fkrp transcript using RNA interference, which showed that the greater reductions of Fkrp mRNA (up to 75%) using dual shRNA cassettes induce overt dystrophic pathology, compared with the lower reductions from using single shRNA cassettes.¹³⁷ However, as shRNA expression by itself can cause toxicity, it remains unclear how much of the phenotype was FKRP specific.¹⁴⁰ Differences in disease severity between the latter two models could also be attributed to laminin α2 localization, which was reduced in FKRP-neo-Y307N mutant muscles but was normal in FKRP-neo-P448L mutants. Interestingly, disease severity may not be correlated with the level of glycosylated αDG expressed, as some expression of glycosylated αDG was detected in FKRP-neo-Y307N muscles by Western blot but not in FKRP-neo-P448L muscles. Taken together, these results suggest that FKRP could have other unknown functions, for example, in regards to laminin α2 localization, and that factors other than glycosylation of αDG may be important for modulating disease severity.

VI. Desmin

Desminopathy is primarily (∼ 80%) a dominantly inherited form of myofibrillar myopathy that leads to cardiac conduction disease, cardiomyopathy, and skeletal muscle weakness.^141,142 The initial symptoms can arise before 10 years of age but are more often reported in the 30s.^141,142 Symptoms usually begin with cardiac conduction disease and progress to cardiomyopathy with severe respiratory distress. There is currently no specific treatment for desminopathies to prevent the progressing cardiomyopathy and deterioration of the diaphragm. Desmin is the primary intermediate filament protein of muscle that connects the sarcomeres to the sarcolemma and organelles.¹⁴³ In the heart, desmin is particularly concentrated in the conduction system, at costameres, and at the desmosome in intercalated disks. Desmin-null mice develop a severe dilated cardiomyopathy from approximately 5 days of age that eventually leads to widespread cardiomyocyte degeneration, fibrosis, and calcification.^144–147 Abnormal mitochondria distribution, proliferation, and swelling are thought to be a central pathologic feature of desmin knockout cardiomyocytes and desmin-related cardiomyopathy.^148–150 The cardiomyocytes are also structurally abnormal in that the sarcomeres do not align and do not attach appropriately to the intercalated disks.^144–147 Desmin knockout mice develop a mild cardiac conduction disease probably owing to the disruption of desmosomes in the conduction system.¹⁵¹ There are two transgenic mouse models that mimic dominantly inherited desminopathy in humans. The first is a 7-amino acid deletion (R173-E179) that leads to chimeric intracellular aggregates, not normally seen in desmin-null mice.¹⁵² The second transgenic mouse is a L345P mutation that significantly alters mitochondrial morphology and function that is associated with altered Ca²⁺ handling.¹⁵³ Together, these transgenic mice suggest that different mutations in desmin can lead to strikingly different phenotypes.

VII. Laminin α2

Laminin α2-deficient mouse models have significantly expanded our understanding of congenital MD type 1A (MDC1A). Models derive from the dystrophic dy/dy mouse line whose pathology originates from mutations in the LAMA2 gene.^154,155 Differences in site and type of mutation determines residual laminin α2 expression, and by extension, disease severity.^156–158 The dy/dy mouse possesses a moderate fibrotic and dystrophic phenotype, coincident with reduced laminin α2 expression.^156,159 The mutation responsible for the dy/dy phenotype is unknown; however, additional mouse models with targeted mutation (dy^2J/dy^2J) or removal (dy^W/dy^W) of the LAMA2 N-terminal domain show minor and severe dystrophic phenotypes, respectively.^156,160,161 Importantly, the severe dystrophic phenotype observed in dy^W/dy^W closely resembles that of dy^3K/dy^3K mice where laminin α2 expression is completely abolished.^156,162 While the secondary mechanisms governing MDC1A pathogenesis remain unclear, these models underscore the connection between laminin α2 expression and disease severity and will likely prove invaluable in the design of novel therapeutic strategies.

A congenital MD was found in a mixed breed dog that had an absence of the laminin isoform α2, which is predominant in, but not specific to, skeletal muscle.¹⁶³ Mutations causing laminin α2 deficiency result in central nervous system defects in humans and peripheral nerve defects in dy mice. It will be interesting to see if similar abnormalities are found in laminin α2-deficient dogs.

VIII. Collagen VI

Collagens are secreted triple-helical extracellular matrix proteins that maintain integrity across many tissues.^164,165 Collagen VI forms a heterotrimer composed of three alpha chains encoded by the COL6A1, COL6A2, and COL6A3 genes. The microfibrils generated upon assembly of collagen VI molecules have been implicated in the maintenance of muscle function and provide an important link between muscle cells and the surrounding matrix through a variety of molecular interactions.^166–168 Mutations in the three collagen VI alpha chains give rise to autosomal dominant Bethlem myopathy (BM)^169–172 as well as the more severe usually autosomal recessive Ullrich congenital muscular dystrophy (UCMD).^173,174 BM is characterized by later onset, proximal muscle weakness with joint contractures, especially of the elbow, finger, and ankle, and slow progression, while UCMD typically manifests early with rapid progression of symptoms that include muscle weakness and hypotonia, joint contractures, and hyperelastic distal joints. Respiratory failure is common in later stages of UCMD and can result in early death if untreated. In both BM and UCMD, intelligence is normal and serum CK levels are normal or slightly elevated.

Investigation of collagen function led to generation of collagen VI-deficient mice through targeted disruption of the col6a1 gene, thus abolishing assembly of the triple helix.¹⁷⁵ The resulting phenotype strongly resembled BM, and these mice became a useful animal model for studying the pathophysiology of both BM and UCMD.¹⁶⁷ Histology and Evans blue dye staining revealed early onset focal necrosis, fiber size variation, and mild central nucleation, all indicators of skeletal muscle pathology. Locomotor performance is not severely affected, though col6a1-deficient mice exhibit muscle weakness, and the condition progresses very slowly, similar to BM. However, the diaphragm is more severely affected than other muscle groups, similar to UCMD.^167,176,177 Col6a1-deficient mice also reflect the dominant character of BM, as heterozygotes show signs of myopathy, though with milder defects.¹⁷⁵

The mouse model of collagen VI deficiencies has revealed important mechanistic insights into the pathophysiology of BM and UCMD, including the involvement of mitochondrial dysfunction and apoptosis,^167,176 but the relationship between these abnormalities and collagen mutations is still poorly understood. Nevertheless, col6a1-deficient mice have allowed testing of specific therapeutic interventions such as cyclosporine A and other pharmacological treatments to target the mitochondrial permeability transition pore.^167,178 While the severity of the deficits observed in UCMD is not captured by these mice, zebrafish models have been reported with early onset, severe myopathy that could be useful for initial screening of novel therapies, and further investigating the molecular etiology of both BM and UCMD.¹⁷⁹

IX. D4Z4 Repeats and FSHD

Facioscapulohumeral muscular dystrophy (FSHD) is caused by macrosatellite DNA contraction in the D4Z4 region of telomeric 4q35.¹⁸⁰ The molecular events following DNA contraction include epigenetic events resulting in chromatin hypomethylation, chromatin relaxation, and expression of a highly conserved double homeobox retrogene, DUX4, embedded within the repeats.¹⁸¹ Recent studies suggest that disease arises only when DUX4 expression occurs in the context of an intact polyadenylation site following the last repeat on the pathogenic alleles and presents a unifying model for FSHD disease mechanism.^182,183

Controversy surrounds the involvement of another gene proximal to 4q35, FRG1. Transgenic mouse studies show that overexpression of FRG1 causes an MD similar to FSHD.¹⁸⁴ Disease severity increases with an increase in FRG1 expression. To date, this is the only model that recapitulates some of the disease features, such as a degenerative muscle phenotype more pronounced in muscles similarly affected in FSHD patients, increased central nuclei, abnormal fiber morphology with increased cross-sectional area, decreased muscle size, kyphosis, and weakness.

Attempts to produce a transgenic mouse line expressing DUX4 have not been successful, but engineering a mouse line that expresses a low level of DUX4 in adult mice to recapitulate DUX4 levels in FSHD patient muscle may prove a difficult task. DUX4 expression in muscle cells in culture results in cell death accompanied by an increased sensitivity to oxidative stress which supports the notion that DUX4 plays a role in early development but is inactivated in the adult. However, because it is generally agreed that overexpression of a gene or genes is the cause of FSHD, therapy development is now underway directed toward eliminating toxicity by targeting mRNA for destruction. Muscle structural and histological changes associated with FRG1 overexpression in the FRG1 transgenic mouse were improved with delivery of constitutively expressed RNA hairpins to direct RNA interference.^140,185 More importantly, functional rescue of muscles, as measured by endurance in treadmill running studies, was possible with AAV6 short hairpin RNA intravascular systemic delivery after the onset of disease.¹⁴⁰ These studies serve as a proof-of-principle for therapeutic antisense approaches for FSHD and other dominant muscle diseases.

X. DMPK and Myotonic Dystrophy

Myotonic dystrophy (DM1) is caused by a microsatellite CTG repeat expansion in the 3′UTR of the dystrophia myotonica protein kinase gene (DMPK).¹⁸⁶ Transcripts with expanded repeats accumulate in the nuclei of cells and alter the availability of proteins involved in the regulation of alternative splicing during development.^187,188 Important protein families for developmental splicing regulation that are affected by the DMPK mRNA repeat expansions are the muscleblind-like (MBNL) and CUGBP/Elav-like family (CELF) proteins.^187,189,190 Disruption of splicing patterns to produce embryonic transcript profiles for a host of unrelated mRNAs results in a disease phenotype with seemingly unrelated features, such as muscle stiffness (myotonia), hyperinsulinemia, gastrointestinal insufficiencies, and cognitive impairment.¹⁸⁶

Approximately 20 transgenic mouse lines have been generated to dissect the molecular pathology resulting from the toxic gain-of-function mRNA expression and to produce models for testing potential therapies for DM1 (reviewed in detail^191a). The models best suited for therapy testing are based on either expressing the CTG repeat expansion or altering the levels of MBNL1 and CUGBP1. One of the most informative and useful transgenic lines, the HSA^LR line, was generated from expression of CTG₂₅₀ from the human α-skeletal actin gene placed in the HSA 3′UTR.^191b The HSA^LR mice display profound myotonia and subsequent changes in MBNL levels that lead to altered splicing profiles. Similar to the human disease, MBNL1 can be found in nuclear accumulations of the protein with the CUG₂₅₀ repeat HSA mRNA. Notably, CUGBP1 levels are not affected, and the muscle-wasting phenotype of DM1 is not reproduced in this transgenic mouse line. Additionally, expression of HSA^LR is limited to muscle and therefore cannot contribute to the effects in multiple organ systems associated with the human condition. Regardless of the restriction of expression of the HSA^LR mRNA in muscle, the HSA^LR mouse has been used for testing antisense therapeutic approaches and has resulted in studies showing elimination of myotonia and reversal of splicing defects with intramuscular delivery of AOs targeting the CUG repeats.¹⁹²

Another notable transgenic line contains a large fragment of the human DMPK locus with an unstable expanded CTG repeat that has become larger with increasing generations to reach 1000–1800 repeats.¹⁹³ The DM500 mice display multisystemic attributes of human DM1, including splicing defects in the muscle, heart, CNS, myotonia, muscle weakness, glucose metabolism defects, and tau distribution abnormalities in the brain. The requirement for homozygosity in these mice suggests that the level of mutant DMPK mRNA expression is below a threshold necessary for pronounced disease and makes these mice expensive and difficult for testing potential therapies.

A third model allowed conditional tissue-specific expression of a 960 repeat expansion in the context of the last exon of DMPK where the expansion normally is found.¹⁹⁴ These mice recapitulate all muscle features of DM1, including muscle wasting, highlighting the importance of the chromosomal context for a more complete disease phenocopy. Other models focus on manipulation of the levels of the MBNL and CLEF protein family members and have limited potential for therapeutic testing related to manipulation of their expression levels. MBNL1 knockout mice recapitulate the splicing defects and myotonia observed with toxic repeat expression.¹⁹⁵ CUGBP1 overexpressing mice display a severe muscle-wasting phenotype not seen in repeat expansion-based models lacking the DMPK chromosomal context, which correlates CUGBP1 upregulation with a muscle-wasting phenotype.¹⁹⁶ Despite the numbers of transgenic mice made to reproduce the many features of DM1, there remain only two lines that have been used for testing potential therapies: the HSA^LR model and the DM500 model (500 repeats in large chrosomal fragment from the DMPK locus). This situation highlights the difficulties often associated with attempts to recapitulate human disease in the mouse with gene insertion, especially related to unstable repetitive and ultimately toxic DNA.

XI. Conclusions

The large and growing number of animal models available to study MD has proven to be an indispensible resource for understanding the pathophysiology of these disorders and for testing experimental approaches to therapy.¹⁹⁷ However, the genes for a number of forms of MD have not yet been identified and there do not yet exist corresponding animal models, and even those genes that have been found have not all been mutated in animals. Developing these newer models is an important goal, especially for the rare MDs where few patient samples are available. In such cases, therapies being developed for other disorders could potentially be quickly tested in animal models for safety prior to therapeutically oriented clinical trials.