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In a forward genetic approach to identify novel genes for congenital muscle diseases, a zebrafish mutant, designated patchytail, was identified that exhibits degenerating muscle fibers with impaired motility behavior. Genetic mapping identified a genomic locus containing the zebrafish ortholog of the dystroglycan gene (DAG1). Patchytail fish contain a point mutation (c.1700T>A) in dag1, resulting in a missense change p.V567D. This change is associated with reduced transcripts and a complete absence of protein. The absence of α-dystroglycan and β-dystroglycan caused destabilization of dystroglycan complex, resulting in membrane damages. Membrane damage was localized on the extracellular matrix at myosepta as well as basement membrane between adjacent myofibers. These studies also identified structural abnormalities in triads at 3 days post fertilization (dpf) of dystroglycan-deficient muscles, significantly preceding sarcolemmal damage that becomes evident at 7 dpf. Immunofluorescence studies identified a subpopulation of dystroglycan that is expressed at t-tubules in normal skeletal muscles. In dag1-mutated fish, smaller and irregular-shaped t-tubule vesicles, as well as highly disorganized terminal cisternae of sarcoplasmic reticulum, were common. In addition to skeletal muscle defects, dag1-mutated fish have brain abnormalities and ocular defects in posterior as well as anterior chambers. These phenotypes of dystroglycan-deficient fish are highly reminiscent of the phenotypes observed in the human conditions muscle–eye–brain disease and Walker–Warburg syndrome. This animal model will provide unique opportunities in the understanding of biological functions of dystroglycan in a wide range of dystroglycanopathies, as disruption of this gene in higher vertebrates results in early embryonic lethality.
Muscular dystrophies are a diverse group of hereditary genetic diseases characterized by progressive weakness and degeneration of skeletal muscles. The membranes of the fibers are fragile and extensive damage occurs leading to necrosis and muscle wasting (1,2). The functioning of skeletal muscle requires that the intracellular sarcomeric proteins stay in association with the membrane and extracellular matrix (ECM). The dystrophin–glycoprotein complex (DGC) is a multimeric protein complex that provides a strong mechanical link from the intracellular cytoskeleton to the ECM (3,4). The components of DGC include transmembrane, cytoplasmic, and extracellular proteins and mutations in most of the members have been identified as the cause of common forms of muscular dystrophies (5–7).
Dystroglycan is an integral membrane component of the DGC, linking dystrophin in the intracellular cytoskeleton to proteins in the ECM (3). The dystroglycan gene (DAG1) is highly conserved throughout evolution and is found as a single copy gene in most vertebrates, including zebrafish (Danio rerio), although several other branches of teleost fish have gene duplications resulting in two copies of the gene (8). Dystroglycan is expressed as a precursor protein, and post-translational processing of the protein through proteolysis cleavage at Ser654 yields two mature proteins, α-dystroglycan and β-dystroglycan (9). α-Dystroglycan is an extracellular protein that binds laminin α2 in the ECM, whereas β-dystroglycan is an integral membrane glycoprotein that connects intracellularly to dystrophin (which binds to the actin cytoskeleton) and extracellularly to α-dystroglycan. In addition to binding the core components of the DGC, dystroglycans are also implicated in signaling processes by binding with proteins involved in signaling pathways (10,11).
Post-translational modification of dystroglycan by glycosylation is required for its function as loss of these modifications results in disruption of dystroglycan–ligand interactions (4). Although genetic mutations of dystroglycan remain to be identified in human patients, recessive mutations in at least six genes resulting in defective glycosylation of dystroglycan lead to genetic forms of muscular dystrophy (5,12–17). Deficiency in dystroglycan glycosylation is common to muscular dystrophies such as Walker–Warburg syndrome (WWS), muscle–eye–brain disease (MEB), Fukuyama congenital muscular dystrophy, congenital muscular dystrophy 1C and 1D and limb girdle muscular dystrophy (18–24), constituting a group of neuromuscular disorders termed ‘dystroglycanopathies’. In several of these dystrophies, in addition to defective glycosylation, a reduction in dystroglycan is also observed at the protein level (22,25).
To understand the molecular pathways and pathogenesis of muscular dystrophies associated with dystroglycan function, a number of animal models have been developed. Dystroglycan and most of the members of DGC are highly conserved in vertebrates and, therefore, offer a wide range of possible animal models to study pathophysiology of the disease. Dystroglycan null mice are embryonic lethal due to very early defects in basement membrane assembly (26). The disruption of Reichert's membrane, a structure that is specific to rodent development, is the primary cause of embryonic lethality in mice. Skeletal muscle-specific ablation of dystroglycan using CMK-Cre mice produced a relatively mild dystrophic phenotype as satellite cells were still able to express dystroglycan and resulted in regenerating muscle fibers (27). Reduced expression of α-dystroglycan is also seen to be associated with dystrophy in Sphynx and Devon Rex cats (28). Transient knockdown of dystroglycan by antisense morpholinos in zebrafish leads to disorganized muscles and apoptotic as well as necrotic cells (29). Drosophila also have a dystroglycan ortholog (Dg) that can undergo glycosylation in vitro (30). However, the enzymes that glycosylate dystroglycan in vertebrates are absent in Drosophila suggesting that post-translational modifications, and therefore its functions, are unlikely to be analogous to that in humans.
In recent years, zebrafish have emerged as a powerful genetic tool to study muscle diseases. All of the dystroglycanopathy genes are highly conserved between mammals and zebrafish (31). Forward as well as reverse genetic approaches employing zebrafish have led to establishing models of several skeletal muscle diseases (32–35). To identify genes responsible for skeletal muscle disorders, an ENU (N-ethyl-N-nitrosourea) screen was performed that identified a group of mutants showing dystrophic and myopathic phenotypes. A mutant, patchytail, was identified in this screen that showed impaired locomotion behavior and dystrophic muscles as identified by birefringence assay. Here we describe genetic and biochemical studies demonstrating a primary defect of dystroglycan in these fish, and show that congenital absence of this protein leads to diverse muscle, ocular and central nervous system (CNS) defects. Thus, the patchytail fish is a good model for early developmental defects in WWS, MEB and other human diseases involving defective glycosylation of dystroglycan.
In an attempt to identify new genes in congenital skeletal muscle diseases, an ENU F2 mutagenesis screen for homozygous recessive mutations was performed in zebrafish (36). Zebrafish embryos with skeletal muscle abnormalities were identified using a birefringence assay that involves examination of axial skeletal muscles of live zebrafish embryos using polarized filter microscopy (37,38). Wild-type embryos from 3 days post fertilization (dpf) onwards display highly birefringent skeletal muscles due to the ordered array of myofilaments (Fig. 1A and B). Skeletal muscle mutants identified with this screen, which displayed reduced or patchy birefringence compared with wild-type embryos, were selected for subsequent studies. One of the mutants, designated patchytail, displayed patchy birefringence indicative of areas of muscle degeneration in somites of homozygotes. During earlier development, the formation of muscle fibers in patchytail was indistinguishable from wild-type embryos. The first signs of muscle degeneration in patchytail were observed at 3–4 dpf and were identified as dark areas in somites under polarized light when compared with wild-type (Fig. 1B and D). In patchytail fish, muscle degeneration is initially restricted to a very few posterior somites at this stage (4 dpf). This is a much milder phenotype in comparison with other dystrophic fish models, such as sapje (dystrophin deficient) (Fig. 1E and F) or candyfloss (‘caf’ or laminin α2 deficient) fish (Fig. 1G and H) where generalized muscle degeneration was seen in most somites by 2–3 dpf (Table 1) (34,35). Muscle degeneration in patchytail progressed rapidly after 4 dpf, and most of the somites exhibited muscle degeneration by 7 dpf (Fig. 1I and J). By this stage, skeletal muscle disorganization was seen randomly distributed in all somites in the trunk. Muscle degeneration in patchytail appeared to be restricted to skeletal muscles. There was no evidence for any morphological abnormalities of the heart, cardiac enlargement or edema of the cardiac sac through 7 dpf. Furthermore, cardiac function appeared to be unaffected as observed by similar heart rates in mutant in comparison with wild-type embryos. A large number of mutant embryos were raised (~300), and all homozygotes died by 8–10 dpf; however, in the heterozygous state, these fish were fully viable, fertile and apparently unaffected.
Mutant patchytail embryos exhibited a defect in motor activity starting early in development. In developing zebrafish, muscle activity results in hatching from their chorion ~2–2.5 dpf. Typically 95 ± 3% (P < 0.005) of wild-type embryos were hatched from their chorions by 2.5 dpf. In contrast, only 85 ± 4% (P < 0.005) of mutant embryos hatched by 2.5 dpf, suggesting a mild muscle weakness during early development.
By 7 dpf, mutant fish exhibited significantly slower swimming than wild-type controls. The swimming phenotype was examined by video microscopy with a touch-evoked escape behavior assay. Typically, wild-type 7 dpf larvae respond to tactile stimuli with a rapid and vigorous escape contraction, followed by swimming, which often resulted in larvae rapidly swimming out of the field of view (Fig. 2A–D). In contrast, patchytail embryos displayed weak escape contractions, followed by reduced swimming that often failed to propel the larvae more than a few lengths (Fig. 2E–J and Supplementary Material, Movies 1 and 2) (wild-type at 7 dpf: 6.2 ± 0.4 cm/0.1 s, n = 10; mutant at 7 dpf: 0.75 ± 0.08 cm/0.1 s, n = 10; Student's t-test, P < 0.001). A highly diminished touch-evoked escape behavior at 7 dpf is indicative of impaired motor function and overall skeletal muscle weakness. Together, these data suggest only mild normal muscle dysfunction during early patchytail development followed by progressive muscle degeneration by 7 dpf suggestive of a dystrophic phenotype.
To identify the mutation underlying the patchytail dystrophic phenotype, genetic mapping was performed. Pooled DNA of 40 patchytail and 40 control siblings of 5 dpf embryos, respectively, were tested with 288 simple sequence length polymorphism markers resulting in linkage of the mutation to chromosome 22. Using standard bulk segregant analysis, a first-pass map position for the patchytail locus was established to markers z6850 (29.1 cM), z9084 (56 cM), z28677 (59.3 cM). Using a fine mapping strategy, this region was further refined to flanking markers z21243 (43.3 cM) and z9084 (55.9 cM), approximating a genomic region of 13 cM (Fig. 3A). As the phenotype observed in patchytail fish suggests a dystrophic process in skeletal muscles, an analysis of this chromosomal area for skeletal muscle-specific genes resulted in identification of the dystroglycan gene (dag1) as a candidate. Sequencing of dag1 revealed a point mutation c.1700 T>A in exon 3 that segregated with the mutant phenotype (Fig. 3B).
The point mutation identified in dag1-mutated fish (c.1700T>A) results in a missense amino acid change of valine (V) to aspartic acid (D) (p.V567D). This change is present in the C-terminal domain of α-dystroglycan that is required for DGC assembly by interacting with the C-terminal domain of β-dystroglycan (Fig. 3C). Computational predictions for the effect of this missense change on protein were evaluated using the Polyphen-2, SIFT and PMut programs. Polyphen-2 predicted this mutation to be ‘probably damaging’ (score 0.969, sensitivity, 0.70, specificity 0.94). Mutation at 567 from V to D was predicted to be not tolerated and affecting protein function with a score of 0.00 by SIFT. Amino acids with probabilities <0.05 are predicted to be deleterious by SIFT. Finally, the pMut program predicted V567D change to be pathological (NN output 0.9055, reliability 8).
The effect of this point mutation in patchytail fish was experimentally evaluated at the RNA and protein levels. RT–PCR showed decreased gel band intensity of dag1 transcripts in patchytail mutant embryos in comparison with the wild-type controls, suggesting that dag1 mRNA levels were likely to be reduced in mutant fish (Fig. 3D). To assess the effect of mutation on α- and β-dystroglycan proteins, western blotting was performed on wild-type as well as patchytail zebrafish embryos. Dystroglycan protein is post-translationally cleaved into α- and β-dystroglycan and expression of both of these polypeptides were examined. Western blot analysis revealed protein products of expected sizes in wild-type embryos. In comparison with the positive controls, no detectable α- or β-dystroglycan expression was seen in mutant embryos (Fig. 3E). Therefore, these studies show that the p.V567D change in dag1 is associated with reduced levels of dag1 mRNA and absence of both α- or β-dystroglycan proteins.
To further validate that the dystrophic phenotype observed in patchytail fish is a result of dystroglycan deficiency, rescue studies were performed. This involved the rescue of mutant phenotype by overexpression of zebrafish dag1 mRNA in embryos. One cell embryos obtained from a cross between two heterozygote patchytail fish were injected with wild-type zebrafish dag1 mRNA. In non-injected clutches, 21.5 ± 2.8% of the embryos showed the mutant phenotype and 78.5 ± 2.8% showed normal phenotype at 4 dpf as seen by birefringence assay. Embryos injected with dag1 mRNA showed a reduction in the mutant phenotype to 10 ± 2.65% of embryos with 90 ± 2.65% of embryos exhibiting normal phenotype (Fig. 3F). Rescue of the mutant phenotype on overexpression of dystroglycan mRNA strongly indicates that the dystrophic phenotype in patchytail fish is a result of dag1 mutation.
Members of the DGC complex are required not only for maintaining stability of the complex but also for the protein stability of their interacting partners. Therefore, the effect of the lack of dystroglycan on stability of other members of DGC complex was evaluated by studying their expression in dag1-mutated fish by immunofluorescence at 7 dpf. In zebrafish embryos, like human embryos, members of the DGC complex are expressed initially at the peripheral ends of myofibers known as myotendenous junctions or myosepta. Immunofluorescence using α- and β-dystroglycan showed that in wild-type embryos these proteins are mainly localized at myosepta in skeletal muscles during embryonic development (Fig. 4A and C). In dag1-mutated embryos, however, a complete absence of α-dystroglycan and β-dystroglycan proteins was observed (Fig. 4B and D). Expression of laminin α2 was observed at the ECM at myosepta between adjacent somites in wild-type embryos. In dystroglycan-deficient embryos reduced expression of laminin α2 was observed compared with wild-type. Moreover, the area of laminin α2 expression at myosepta appeared widened in comparison with wild-type myosepta (Fig. 4E and F, arrow). Dystrophin expression was also detected at the myosepta in wild-type embryos. The expression of dystrophin in dystroglycan-deficient fish was seen to be highly reduced (Fig. 4G and H). These studies demonstrate that dystroglycan deficiency results in destabilization of their interacting partners in both extracellular as well as the intra-cellular side of the myoseptum.
To evaluate the loss of dag1 on structure of affected skeletal muscles, toluidine blue-stained longitudinal sections of wild-type and dystroglycan-deficient fish were examined at 7 dpf (Fig. 5A–D). Myofibers in both wild-type and mutant fish looked well organized. However, the myosepta in mutant fish appeared to be thickened in width compared with wild-type siblings. Higher magnification views revealed gaps at myosepta between adjacent somites (Fig. 5D, arrow). Dystroglycan-deficient fish also showed gaps between adjacent myofibers, suggesting fiber detachment at ECM (Fig. 5D, arrowhead).
Chimeric mice deficient for dystroglycan exhibit a neuromuscular junction defect with fewer and fragmented nerve synapses in the affected muscles (39). Therefore, the effect of dystroglycan deficiency on zebrafish neuromuscular junctions was also evaluated. Whole-mount labeling for postsynaptic acetylcholine receptors using fluorescent-conjugated α-bungarotoxin was performed for dystroglycan-deficient as well as wild-type embryos (Fig. 5E and F). Acetylcholine receptors were found to be localized to myofiber ends adjacent to myoseptum as well as with in the somites in wild-type embryos (Fig. 5E). Similarly, dystroglycan-deficient embryos showed well-developed postsynaptic receptor clusters. There were no discernable differences between wild-type and dystroglycan-deficient fish in the number and size of clusters, which suggests that dystroglycan may not be required for neuromuscular junction formation in zebrafish muscles.
To visualize the ultrastructure of skeletal muscles, transmission electron microscopy was performed. Dystroglycan-deficient fish display a mild dystrophic phenotype during early development (3 dpf) followed by severe impairment of locomotive behavior by 7 dpf. Therefore, to understand the pathological progression of muscle degeneration, ultrastructure of dag1-mutated fish was examined at these two time points during zebrafish development. Early in development, myospeta, in most of the somites of the dystroglycan-deficient fish, were organized similar to wild-type siblings (Fig. 6A and B). In some somites of the mutant fish, initiation of tearing, identified by the lack of electron dense areas in a myoseptum, was observed at this stage (Fig. 6B, arrow). Despite this minor tearing, myofibers were largely intact in the mutant fish (Fig. 6B, arrowhead). By 7 dpf, ultrastructure of dystroglycan-deficient fish showed highly distorted and irregular-shaped myospetal boundaries in most of the somites (Fig. 6C and D). The myosepta of mutant embryos appeared to be fragmented with regions of separated ECM (Fig. 6D, arrowhead).
Around 7 dpf during normal development, expression of DGC components becomes prominent in the sarcolemma of myofibers (40). To investigate whether the loss of dystroglycan affects sarcolemmal integrity, ultrastructure of sarcolemma of wild-type and dag1-mutated embryos were evaluated at 7 dpf. The cross-sectional view of wild-type zebrafish muscle fibers showed well-organized sarcolemmal boundaries between adjacent myofibers (Fig. 6E). However, the sarcolemmal membranes in dag1-mutated embryos appeared to be grossly disorganized when compared with wild-type muscles (Fig. 6F). High magnification examination revealed widening of the area between myofibers of two adjacent fibers on the ECM side (Fig. 6H, arrows). However, no such gaps were observed between wild-type myofibers (Fig. 6G, arrows). The sarcolemmal integrity was evaluated in vivo using Evans blue dye that labels cells with damaged plasma membranes. No uptake of dye was observed in either wild-type or dystroglycan-deficient fish, suggesting that the sarcolemma remains largely intact in dag1-mutant fish. These data demonstrate that severe dystrophy observed in dystroglycan-deficient fish is due to massive detachment in extracellular junctions at myoseptum as well as disruption of extracellular integrity at the sarcolemmal membranes.
Further analysis of myofiber ultrastructure in dystroglycan-deficient fish showed normal organization of the contractile apparatus with no necrotic fibers or apoptotic nuclei through 7 dpf (Fig. 7A–D). While the contractile apparatus was structurally normal appearing in mutant fish, higher magnification views revealed that t-tubules in dag1-deficient fish were smaller and disorganized in comparison with wild-type controls even during earlier stages of development (3 dpf) (Fig. 7B, arrow). Disorganized t-tubules were very rare in wild-type controls, but were observed even in somites of patchytail fish where the myoseptum was seen to be intact, suggesting that these effects are independent of each other. No significant changes in sarcoplasmic reticulum were observed at 3 dpf. These observations suggest that t-tubule disorganization is an early event in the pathogenesis of the disease. During the larval stages at 7 dpf, most of the myofibers continued to have highly disorganized t-tubules (Fig. 7D, arrow). Additionally, terminal cisternae of sarcoplasmic reticulum, which are localized adjacent to t-tubules, also appeared to be collapsed in most of the myofibers in comparison with control fish (Fig. 7D, arrowhead). No significant abnormality was detected in the longitudinal vesicles of the sarcoplasmic reticulum.
Thus far, expression of dystroglycan has been reported at the sarcolemma in skeletal muscles cells. To understand whether the t-tubule defects are a direct result of loss of function of dystroglycan, indirect immunofluorescence was performed to evaluate the subcellular localization of dystroglycan in developing zebrafish muscles. In wild-type fish, α- and β-dystroglycan are expressed both at the sarcolemma and within myofibers in a striated pattern at 7 dpf (Fig. 7E and F). Co-immunostaining for dystroglycan and the t-tubule marker, Dhpr, showed that α- and β-dystroglycan co-localize with Dhpr in wild-type myofibers (Fig. 7G and I). As expected, no expression of α- and β-dystroglycan was observed in myofibers from patchytail fish. Interestingly, Dhpr levels appeared to be reduced significantly in the dag1-mutant fish (Fig. 7H and J). Co-immunostaining for ryanodine receptors, Ryr1, and Dhpr showed that, as expected, they co-localized at t-tubules and sarcoplasmic reticulum junction (i.e. the triads) in wild-type fish. While moderate levels of Ryr1 expression were detected in dystroglycan-deficient fish, expression of Dhpr was again highly diminished (Fig. 7K and L). These data suggest that, in addition to expression at the sarcolemma, a significant fraction of dystroglycan is also localized at the t-tubules in wild-type fish where it may play a role in organizing or stabilizing Dhpr calcium channels.
Dystroglycan is widely expressed in different organs in addition to skeletal muscles in mammals. To study the expression of dystroglycan in different organs in zebrafish, whole-mount immunofluorescence was performed. Immunofluorescence studies showed that besides expression at the myosepta in skeletal muscles, dystroglycan is also expressed in the eyes, CNS and heart in wild-type fish (Fig. 8A). In dag1-mutated fish, a complete absence of dystroglycan protein was observed in all organs (Fig. 8B). Owing to strong expression in the eyes and CNS and the involvement of these organs in a wide range of dystroglycanopathies, the effect of dystroglycan deficiency on these organs was evaluated in patchytail fish.
Upon gross examination, the eyes of 5 dpf patchytail embryos appeared to be normal (Fig. 8C and D); however, histological examination revealed striking abnormalities (Fig. 8E and F). At this stage, wild-type eyes are well organized into photoreceptor, inner and outer plexiform layers, ganglion cell layer, lens, and well-developed cornea. In dystroglycan-deficient fish, differentiated cell layers were also present in the eyes. However, in the posterior chamber, cells within the ganglion layers were loosely packed in comparison with wild-type fish and there were gaps present between cells in the ganglion layer in dystroglycan-deficient fish (Fig. 8F, arrow).
The anterior chamber of the mutant eyes showed the most drastic changes relative to wild-type fish. By 5 dpf, wild-type fish lens fibers have differentiated, lost their nuclei and become crystalline. The lens in dag1-mutated fish, however, was cellular with several inclusion bodies present (Fig. 8F, asterisk). The cornea of the eye was absent in dag1-mutated fish suggesting either it was not formed or degenerated due to dystroglycan deficiency.
The effect of dystroglycan loss on CNS development was also studied in patchytail. Histology of control brains showed that by 5 dpf, the CNS is well organized into different chambers; forebrain (telencephalon), midbrain (tectum) and hind brain (cerebellum and myelencephalon) (Fig. 9A). When compared with wild-type brain, all four chambers were seen in dystroglycan-deficient fish as well (Fig. 9B). However, tectal cells in midbrain and granular cell layer in cerebellum appeared to be less organized in dystroglycan-deficient fish when compared with wild-type (Fig. 9C–F, arrows). The mutant brain did not show any hydrocephalus or neuronal heterotopia at this stage of development as reported in dystroglycan-deficient mice.
Dystroglycanopathies are a group of inherited muscular dystrophies that result from hypoglycosylation of α-dystroglycan, thus abolishing its interaction with extracellular laminin and resulting in muscle weakness (5). In the present study, we address the role of dystroglycan in muscle development and homeostasis with the aim of understanding the primary role of dystroglycan in human dystrophies. Using zebrafish as a model system, we found that dystroglycan is not required for muscle formation during early development. At later stages of development, the loss of dystroglycan results in muscle degeneration and impaired locomotion. Additionally, we discovered that deficiency of dystroglycan results in t-tubule defects that precede the damage to membranes. Most importantly, the loss of dystroglycan resulted in extensive tearing of ECM at the myosepta, thus destabilizing myofiber attachments at these somite boundaries.
In dystroglycan-deficient fish, only a mild motor phenotype was seen during earlier stages of zebrafish development. Pathologically, no significant myoseptum damage was seen until 7 dpf in dystroglycan-deficient fish, suggesting that dystroglycan is not required for early embryonic development of zebrafish. The only pathological change observed at earlier stages of development was highly disorganized t-tubules in dystroglycan-deficient fish at 3 dpf. T-tubules are invaginations of plasma membrane that, together with sarcoplasmic reticulum, form the triad through which calcium homeostasis and excitation–contraction coupling of muscle fibers occurs (41). Although topologically continuous with each other, the protein and lipid composition of the t-tubules has been shown to be distinct from that of the sarcolemma, suggesting that these membrane fractions are evolved to perform distinct functions (42). Our immunofluorescence studies showed that dystroglycan co-localized with the t-tubule marker Dhpr in wild-type fish, suggesting that the disorganized t-tubules are not a result of secondary effects of dystroglycan deficiency but rather are a primary consequence. Furthermore, the level of immunostaining for the t-tubule marker, Dhpr, was greatly reduced in dag1-mutant fish, while Ryr1 staining was only moderately affected, consistent with our observation that dystroglycan deficiency had an earlier and greater impact on t-tubule morphology than on the sarcoplasmic reticulum. Previous studies have identified the expression of other members of DGC complex in t-tubules (43–47), which is not surprising considering that they are membrane invaginations of the sarcolemma and, therefore, may retain several protein components that are also present on sarcolemma. Dystrophin interacts either directly or indirectly with Dhpr and is believed to regulate channel function that is disrupted in DMD muscles (43). Our data suggest a model whereby dystroglycan may play a key role in this interaction, perhaps by linking dystrophin to DHPR channels and stabilizing their localization in t-tubule membranes. Regardless, abnormal t-tubule morphology in patchytail fish supports the notion that DGC components in t-tubules are required for structural and/or functional properties and suggests that defective excitation–contraction coupling should be considered as a possible contributing factor to the pathogenesis of skeletal muscle weakness in human patients with dystroglycanopathies. Thus, in dystrophic muscles from patients with dystroglycanopathies, additional pathways may be involved in addition to the primary sarcolemmal membrane defects that are considered to be the central underlying mechanism of muscle degeneration in these conditions.
The most striking pathological change in dystroglycan-deficient muscle was the extensive tearing of the myosepta, which developed between 3 and 7 dpf. This degeneration is likely due to the absence of complex formation between laminin α2 and α-dystroglycan that otherwise provides stability to muscle fibers (48,49). While reduced levels of laminin α2 and dystrophin were seen in dystroglycan-deficient fish, the pathology of the disease in patchytail is different than observed in most forms of dystrophies (Table 1). In sapje fish, a lack of dystrophin results in detachment of terminal sarcolemma from myoseptum, leading to a collapse of the contractile apparatus and cell death (34). Softy fish have a mutation in the ECM gene lamb2 and show adhesion failure of myofibers during very early in development (30 hours post fertilization) (50). The pathological changes observed at the myosepta of patchytail fish most closely resemble those seen in caf fish with a lama2 mutation in which myofiber detachment occurs on the extracellular side of the membrane at myosepta due to damage at the ECM rather than at the sarcolemma (35). Thus impaired dystroglycan–laminin interactions are the common factor underlying the pathogenetic mechanisms of these disorders. Previous studies on nonmuscle cells have also suggested that dystroglycan is required for organization of the ECM, as dystroglycan deficiency, and consequent loss of linkage with laminin, leads to disruption of the Reichert's membrane and embryonic lethality in mice (26). Similarly, dystroglycan–ligand interactions are required for the integrity of basement membranes in the brain, cornea and retina of higher vertebrates and disruption of this function results in abnormal development of these tissues in mice and in WWS (51). In several other dystrophic fish models, significant regeneration is seen in the skeletal muscles (40). While most of these strains survive until 3 weeks, no dag1-mutated fish survived more than 10 dpf. As dystroglycan is implicated in muscle regeneration owing to its expression on muscle satellite cells, this severity in phenotype of dag1-mutated fish could also be due to a lack of regenerative capacity of muscle cells. Indeed, no significant increase in regenerating fibers was observed in dystroglycan-deficient fish. Dystroglycan-deficient fish showed indistinguishable cardiac morphology and heart rates in comparison with wild-type fish. This may seem surprising in light of cardiac expression of dystroglycan; however, it is worth noting that human patients with dystroglycanopathy typically do not have cardiac involvement. Nevertheless, we cannot rule out subtle cardiac defects or susceptibility to degenerative changes since dystroglycan-deficient fish die by 8–10 dpf.
Dystroglycan deficiency also caused abnormal development of ganglion, lens and cornea layers in eye in patchytail fish. Similar eye abnormalities have been observed in mice and humans with mutations in several genes that encode proteins involved in glycosylation of dystroglycan (52–54) and mice with epiblast-specific deletion of dystroglycan also develop eye defects that broadly resemble WWS in humans (51,55). Together, these data suggest that eye abnormalities are a direct consequence of dystroglycan deficiency in a wide range of dystroglycanopathies. Chimeric mice deficient in dystroglycan showed disrupted myoneural synapses at neuromuscular junctions (39). The epiblast-specific loss of dystroglycan results in aberrant neuronal migration, lissencephaly and hydrocephalus in the brain (51). Patchytail fish showed only subtle defects in cell organization in tectum and cerebellum at 5 dpf and no significant neuromuscular junction defects. A previous published report has shown no brain abnormalities in a knockdown model of zebrafish dystroglycan, suggesting that dystroglycan function may not be required for brain formation in zebrafish (29). However, further detailed studies of patchytail fish will be able to help understanding the function of dystroglycan in zebrafish brain.
The patchytail zebrafish model provides a unique model to study human dystroglycanopathies as the rapid ex vivo development of this species circumvents early embryonic lethality seen in higher vertebrate models deficient in dystroglycan or enzymes modifying dystroglycan such as fukutin or Pomt1. Specifically, the dystrophic changes in dystroglycan deficiency are due to damage to the ECM at myosepta and between myofibers, thus affecting muscle stability. These results suggest that therapeutics designed to target correcting these pathologies may provide effective treatment for dystroglycanopathies.
Fish were bred and maintained using standard methods as described (56). Wild-type embryos were obtained from Oregan AB line and were staged by hours (h) or days (d) post fertilization at 28.5°C. All animal work was performed with approval from the Children's Hospital Boston Animal Care and Use Committee.
AB males were treated with 3 mm ENU once a week for 3 weeks. The males were then crossed repeatedly to clean out any post meiotic germ cells that were mutagenized. Mutagenized males were then crossed to wild-type AB females and the progeny (F1) were raised. F2 embryos were obtained by setting up at least six pairs of F1 crosses and screened for skeletal muscle defects.
Zebrafish larvae were screened at 3–4 dpf using a birefringence assay. Crosses in which 25 ± 5% of larvae that showed patchy or reduced birefringence of their axial skeletal muscles in polarized light were identified as potential skeletal muscle mutants and these families were selected for further study.
Embryonic locomotor assay was performed as described (57). Briefly, mechanosensory stimuli were delivered to embryo tails using insect pins. Time-lapse images of zebrafish embryos were taken at different time intervals using a Nikon smz1500 microscope with a SPOT camera system.
AB strain dag1-mutated heterozygous zebrafish were out-crossed to wild-type wik to generate polymorphic mapping strains. Low-resolution mapping was done with 40 diploid mutant and 40 diploid wild-type embryos obtained from in-crossing mapping F2 fish. Microsatellite CA markers throughout the genome were used to scan for linkage as described (58).
The potential effect of the V567D substitution was evaluated using the PolyPhen-2 (Polymorphism Phenotyping) (http://genetics.bwh.harvard.edu/pph2/), SIFT (Sorting Intolerant from Tolerant) (http://blocks.fhcrc.org/sift/SIFT.html) and PMut (http://mmb2.pcb.ub.es:8080/PMut/) programs. PolyPhen-2 is based on an empirical set of rules that combine analyses of a pre-built multiple protein sequence alignment with a number of protein structural and functional attributes to estimate the consequence of an amino acid substitution (59,60). Polyphen-2 score is based on the Naïve Bayes posterior probability with larger values reflecting the higher likelihood of a variant to be damaging. SIFT predicts the effect of an amino acid substitution on protein function based on sequence homology and the physical properties of amino acids (61). PMut is a program that uses neural networks trained on known disease-causing mutations and neutral mutations to provide the binary predictions of ‘neutral’ or ‘pathologic’ (62).
RNA was isolated from pools of mutant or wild-type embryos at 5 dpf using RNeasy fibrous tissue mini kit (Qiagen). cDNA was prepared using superscript III first-strand synthesis kit (Invitrogen). Equal concentrations of wild-type and mutant RNA were used for cDNA synthesis. Following RT–PCR, the amplified DNA bands were quantified using Quantity one software (Biorad).
Zebrafish cDNA library was prepared from 5 dpf AB embryos using Superscript III first-strand synthesis system for RT–PCR (Invitrogen). dag1 cDNA was amplified using forward primer: 5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTYYATGCGCAATAAACTCAGAGAGTTC-3′ and reverse primer: 5′-GGGGACCACTTTGTACAAGAAAGCTGGGTYGGGTGGCACGTAAGGGG-3′ and cloned in to PCSdest vector (a gift from Nathan Lawson) using gateway technology. Capped and polyadenylated, sense dag1 transcripts were synthesized by in vitro transcription using mMessage kit (Ambion). mRNA were injected into 1-cell embryos obtained from cross between two heterozygote dag1 fish. Clutches from five different mating pairs (100–150 embryos per clutch) were injected with dag1 mRNA for rescue experiments.
Fish embryos and larvae were anesthetized and fixed overnight in 4% paraformaldehyde in phosphate-buffer saline (PBS) at 4°C. Embryos were washed with 1× PBS twice and incubated successively in 10 and 20% sucrose solution in 1× PBS till embryos sank. Embryos were embedded in CRYO OCT compound (Fisher) and frozen in 2-methyl butane in liquid nitrogen. Seven micrometer-thick frozen sections were cut using Leica cryostat.
Frozen tissue sections were fixed in methanol at −20°C and washed with tris-buffered saline Tween-20 (TBST) (0.1% Tween 20) followed by blocking in 5% goat serum in TBST for 1 h at room temperature. Sections were incubated with primary antibodies overnight at 4°C followed by incubation with secondary antibodies at room temperature. Imaging was performed using a Nikon eclipse 90i microscope. Whole-mount immunofluorescence was performed as described previously (56). Primary antibodies used in this study were α-dystroglycan (VIA4-1, Millipore), β-dystroglycan (Novocastra), laminin α2 (Sigma), β-actin (Sigma) and dystrophin (Sigma). Nuclear staining was done using DAPI. Secondary antibodies were purchased from Jackson ImmunoResearch.
Evans blue dye staining was performed essentially as described previously (34). Evans blue dye was injected at 0.1 mg/ml into the pre-cardiac sinus of anesthetized embryos at 3 dpf. Embryos were examined and photographed 4–6 h later.
Embryos were fixed in 4% paraformaldehyde for 2 h at room temperature and dehydrated in 100% methanol overnight (−20°C). Embryos were subsequently rehydrated and permeabilized in 1× TBST (1% tritonX-100) for 2 h at room temperature. Blocking was performed with 5% goat serum in TBST for 1 h at room temperature followed by incubation with fluorescent conjugated α-bungarotoxin (5 µg/ml) for 30 min at room temperature. Embryos were washed several times in TBST and mounted in 70% glycerol and visualized using a Nikon eclipse 90i microscope.
One hundred embryos of wild-type or mutant fish at 5 dpf were extracted with 50 mm Tris–HCl buffer containing 150 mm NaCl and 2% Triton X-100. Following centrifugation at 14 000g at 4°C for 20 min, the concentration of protein in supernatants was determined by the Bradford method, using a protein assay kit (Bio-rad).
For the purification of a dystroglycan, wheat germ agglutinin (WGA)–agarose beads (Vector Labs) were added to extracts containing equal amounts (500 mg) of total protein and incubated overnight at 4°C followed by elution from WGA beads with sodium dodecyl sulfate sample buffer (Invitrogen). Proteins were separated by electrophoresis on 4–12% gradient Tris–glycine gels (Invitrogen) and transferred onto polyvinylidene difluoride membranes (Invitrogen). Membranes were blocked in PBS containing 2% casein, 0.1% Tween 20, and were incubated with anti-α-dystroglycan (1:50, VIA4-1, Millipore), anti-β-dystroglycan (1:100, Novocastra) or anti-β-actin (1:1000, Sigma). After washing, the membranes were incubated with horseradish peroxidase secondary antibody (anti-mouse IgG, 1:2000, Jackson Immunoresearch). Proteins were detected using a western blotting detection kit (Millipore).
Zebrafish embryos were fixed in formaldehyde–glutaraldehyde–picric acid in cacodylate buffer overnight at 4°C followed by osmication and uranyl acetate staining. Subsequently, embryos were dehydrated in a series of ethanol washes and finally embedded in Taab epon (Marivac Ltd., Nova Scotia, Canada). Ninety-five nanometer sections were cut with a Leica ultracut microtome, picked up on 100 m formvar-coated Cu grids and stained with 0.2% lead citrate. Sections were viewed and imaged under the Philips Tecnai BioTwin Spirit Electron Microscope (Electron Microscopy Core, Harvard Medical School).
This work was supported by National Institute of Health grants from the National Institute of Arthritis and Musculoskeletal and Skin Diseases (R01 AR044345 to A.H.B.); National Institute of Neurological Disorders and Strokes (P50 NS040828 to A.H.B.); by the Lee and Penny Anderson Family Foundation (to A.H.B.); and the William Randolph Hearst Fund (to V.G.). DNA sequencing was performed by the Children's Hospital Boston Genomics Program Molecular Genetics Core supported in part by National Institute of Child Health and Human Development grant (P30HD18655).
We thank Himani Chinnapen for assistance with the ENU mutagenesis screen and Marie Discenza, Amelia Geggel and Ryan Darnall for their help with zebrafish crosses. We thank Chiara Manzini for wonderful discussions regarding zebrafish brain anatomy. Special thanks to Chris Lawrence from the Zebrafish Core facility at Children's Hospital Boston for wonderful help with all aspects of zebrafish husbandry. We thank Louise Trakimas from Electron Microscopy Core at Harvard Medical School for assistance with histology work.
Conflict of Interest statement. None declared.