Zebrafish muscular disease models towards drug discovery

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    10.1517/17460440902835483 2009 Informa UK Ltd ISSN 1746-0441 507All rights reserved: reproduction in whole or in part not permitted

    ZebrafishmusculardiseasemodelstowardsdrugdiscoveryHiromi HirataNagoya University, Graduate School of Science, Furo-cho, Chikusa-ku, Nagoya 464-8602, Japan

    Background: Zebrafish is an amenable vertebrate model useful for the study of development and genetics. Small molecule screenings in zebrafish have successfully identified several drugs that affect developmental process. Objective: This review covers the basics of zebrafish muscle system such as muscle development and muscle defects. It also reviews the potential use of zebrafish for chemical screening with regards to muscle disorders. Conclusion: During embryogenesis, zebrafish start to coil their body by contracting trunk muscles 17 h postfertilization, indicating that a motor circuit and skeletal muscle are functionally developed at early stages. Mutagenesis screens in zebrafish have identified many motility mutants that display morphological or functional defects in the CNS, clustering defects of acetylcholine receptors at the neuromuscular junctions or pathological defects of muscles. Most of the muscular mutants are useful as animal models of human muscle disease such as muscle dystrophy. As zebrafish live in water, pharmacological drugs are easily assayable during development, and thus zebrafish may be used to determine novel drugs that mitigate muscle disease.

    Keywords: development, locomotion, muscle, mutant, zebrafish

    Expert Opin. Drug Discov. (2009) 4(5):507-513

    1. Developmentofskeletalmuscleinzebrafish

    Zebrafish have three distinct cell types of axial musculature; fast-twitch muscle cells (fast muscle), slow-twitch muscle cells (slow muscle) and muscle pioneer cells. The fast and slow muscles are identical to white and red muscles, respec-tively. Muscle pioneers, which are necessary for correct outgrowth of primary motor axons, are a specialized subset of slow muscle cells in fish. Each muscle cell originates from somites but takes a distinct lineage and is finally located at a distinct part of the axial musculature.

    After gastrulation (5 10 h postfertilization [hpf ]), somites are formed from paraxial mesodermal tissue, in other words, from presomitic mesoderm [1,2]. This segmentation process occurs every 30 min in zebrafish from the anterior to the posterior direction on both sides of the neural tube and notochord. Each somite gives rise to myotomes, dermatomes and sclerotomes, which respectively differentiate into muscles, dermis and bones. In parallel with the somitogenesis, hedgehog signals from midline induce medial myotome to differentiate into adaxial cells, which are muscle precursors that give rise to slow muscles and muscle pioneer cells [3,4]. Most adaxial cells migrate radially to the lateral surface to form a single-layer muscle underneath the skin, where they differentiate into slow muscle cells, whereas a subset of the adaxial cells remains at the medial loca-tion, elongates its shape to span from medial to lateral myotome and differentiates into muscle pioneer cells [5-7]. The rest of the myotomal cells differentiate into fast muscle cells.

    In mammalian musculature, slow and fast muscle cells exist as a mosaic pattern. Each muscle cell type can be histologically distinguished by ATPase

    1. Development of skeletal muscle

    in zebrafish

    2. Development of motility

    in zebrafish

    3. Zebrafish motility mutants

    caused by muscle or

    neuromuscular junction defects

    4. Chemical biology in zebrafish

    5. Conclusion

    6. Expert opinion

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    activity of myosin or reductase activity of mitochondria. In fish, in contrast, these two populations are easily distinguishable, because they are not mixed in location. The slow muscle is only a superficial single cell layer of fibers, whereas the rest of the muscle constitutes fast muscle. In slow muscle, a network of electrical coupling to share synaptic currents is important to drive rhythmic swimming, whereas fast muscle generates action potentials to mediate rapid escape behavior [8-10]. It may be beneficial to examine cell type-specific injury and recovery of muscle in zebrafish.

    2. Developmentofmotilityinzebrafish

    Development of neural circuits and muscles in zebrafish is very fast and embryos show three distinct stereotyped behaviors (spontaneous coiling, touch-evoked coiling and swimming) by 36 hpf [11,12]. The earliest locomotion consists of repetitive, spontaneous alternating coiling of the trunk. This simple slow coiling is independent of mechanosensory stimulation and abruptly starts at 17 hpf, reaches a peak frequency of 0.3 1 Hz at 19 hpf and declines to < 0.1 Hz by 26 hpf. After 21 hpf, embryos respond to mechanosen-sory stimulation with fast trunk coils. The initiation of this touch-evoked coiling indicates that neural networks from somatic sensory neurons to motoneurons through interneurons as well as skeletal muscles are functionally connected even before 24 hpf to execute escape behaviors. By 26 hpf, mechanosensory stimulation initiates swimming, which is defined as a forward movement with rhythmic tail flips by at least one body length. The frequency of muscle contractions during swimming increases from 7 Hz at 26 hpf to 30 at 36 hpf, the latter being similar to the frequency of swimming of adult zebrafish [9].

    The escape behavior followed by a mechanosensory stimulation can be divided into several steps from sensory perception to muscle contraction. In zebrafish embryos, two types of mechanosensory neurons perceive touch stimuli. Head and yolk stimulation are transduced by trigeminal sensory neurons, whereas tail stimulation activates RohonBeard mechanosensory neurons in the trunk [13,14]. RohonBeard cells die within 4 days in development and, in larval stage, the function is taken over by the dorsal root ganglia [15]. These sensory neurons project to interneuronal networks located in the spinal cord and hindbrain to produce motor rhythm [16]. This motor pattern alternatively activates motor neurons in each side of the spinal cord. Motor terminals release acetylcholine at the neuromuscular junctions (NMJs) and depolarize the muscle membrane [9,17,18]. Depolarizations of the plasma membrane spread down the transverse-tubules (t-tubules), which are invaginations of the plasma membrane, and cause conformational changes of the dihydropyridine receptor (DHPR), a voltage sensor located in the t-tubule membrane [19]. The DHPRs then trigger opening of ryano-dine receptors (RyR) in the adjacent sarcoplasmic reticulum

    (SR) membrane to allow Ca2+ release from the SR to the cytosol [20]. Increased cytosolic Ca2+ activates troponin C that initiates actin/myosin sliding, thus causing muscle contraction [21]. The cytosolic Ca2+ levels are rapidly decreased by sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA), a calcium pump expressed in the SR of skeletal muscle that enables fast relaxation [22]. As activation of muscle is caused by rhythmic motor outputs, slow and fast bilateral alternation of muscle contractions generates coiling and swimming behavior, respectively.

    3. Zebrafishmotilitymutantscausedbymuscleorneuromuscularjunctiondefects

    Zebrafish is the vertebrate most amenable to forward genetic screens. Generation of zebrafish homozygous mutants was first described in 1981 by Streisinger and his colleague, who developed zebrafish as a genetic model of vertebrates [23-27]. In the early 1990s two large-scale mutagenesis screens were successfully performed in Tbingen, Germany, and Boston, US, and > 4,000 mutants were identified [28,29]. In these screening, 166 behavioral mutants that displayed abnormal touch-evoked swimming were reported [30]. Among them, in 63 mutations, striation of somitic muscle fibers was reduced by birefringence observation, indicating that these mutants have defects in structural arrangement of actin and myosin. In fact, the responsible genes of this class of mutants are structural components of muscle. On the other hand, some other locomotion mutants displaying normal birefringence under polarized light are defective in Ca regulation in muscle or formation of CNS or NMJ.

    Mutations in dystrophin cause embryonic-onset, progressive degeneration of skeletal muscle and impaired locomotion [31,32]. Many muscle fibers in the mutants were detached from the myoseptum, which is an attachment site of myofibrils located at somite boundaries. Similarly, laminin alpha2 mutants display disconnection of muscle fibers [33]. Although these two mutants show similar fiber detachment and reduced muscle birefringence, the morphological defects in dystrophin mutants are caused by sarcolemmal rupture, whereas the muscle atrophies in the laminin mutants are mechanically induced by spontaneous muscle contraction [33]. Another mutant, which is linked to titin locus, also shows sarcomere defects [34,35]. As mutations of the dystrophin, laminin or titin gene in humans are responsible for muscular dystrophy, these zebrafish mutants could serve as animal models for human muscular dystrophy [36-38].

    Some other mutants with reduced birefringence of skeletal muscle also show morphological defects of muscle. In hsp901 mutants, thick filaments composed of myosin pro-teins are absent and the sarcomere structures are defi-cient [39]. The mutant embryos lack both spontaneous and stimulus-evoked muscle contractility. Similarly, unc45b mutations impair skeletal myofibril formation and locomotion [40]. Hsp901 and its cochaperone Unc45b

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    protein colocalize with myosin and promote myosin assembly during myofibrillogenesis [41-43]. Although these genes encoding for molecular chaperons are responsible for muscle defects in fish, the link between the chaperones and muscle disease is not clear in humans. Another mutant named fibrils unbundled, which is one of the oldest mutants screened in the 1980s, also shows structural defects of skeletal myofibrils. However, the responsible gene for this muscle disorganization is still unknown [44,45].

    Some zebrafish mutants that display normal birefringence but show reduced motility are defective in Ca regulation. The DHPR complex in skeletal muscle is composed of voltage-sensing and pore-forming 1S, cytoplasmic b1a and auxiliary 2d1 and g1 subunits [19]. Nonsense mutations in the cacnb1 gene encoding for the b1a subunit impair excitationcontraction coupling, which is essential for Ca transients during muscle contraction, and thus the mutations cause skeletal paralysis [46-49].As mutations in human DHPR components are responsible for periodic paralysis, a congenital muscle weakness [50], zebrafish cacnb1 mutants are useful for physiological analysis of the disease. A mutation that causes abnormal splicing of ryr1b gene, which encodes for RyR1 in skeletal muscle, also shows defects in excitationcontraction coupling that result in slow swimming owing to weak con-tractions of muscle [51]. Morphologically, this zebrafish mutant displays a number of amorphous cores in muscle, which can also be observed in histological sections of muscu-lature from human multi-minicore disease patients [52,53]. As some susceptible individuals carry splicing mutations in human RyR1 gene [54], the zebrafish ryr1b mutants provide a pathological model of the multi-minicore disease. In fact, application of antisense morpholino oligonucleotides that block the aberrant splicing and thus restore the normal mRNA successfully recovers the normal behavior in the zebrafish mutants [51]. In contrast to these DHPR and RyR mutations, which impair Ca release from the SR and lead to weak contractility of muscle, mutations in atp2a1 gene encoding for SERCA, a Ca pump that is responsible for pumping Ca from the cytosol to the SR, cause overcontraction of muscle [55,56]. In the mutant muscles, duration of the Ca transients is more prolonged than normal that in turn results in stiffness of trunk muscles. As ATP2A1 mutations in humans lead to Brody disease, an exercise-induced muscle relaxation disorder [57], zebrafish atp2a1 mutants could be a useful animal model for this condition.

    Zebrafish mutants that have defects in the NMJ also show reduced motility and normal birefringence. In mam-mals, the postsynaptic acetylcholine receptor (AChR) clusters are formed in an agrin-MuSK dependent manner [58]. Agrin, a nerve-derived ligand, binds to low-density lipoprotein receptor 4 on the muscle surface and activates MuSK (muscle-specific kinase), thereby recruiting AChRs and cyto-plasmic anchoring proteins such as rapsyn at the postsynaptic area [59,60]. Zebrafish chrna1/AChR1 mutants and chrnd/AChRd mutants are immotile, because they lack functional

    AChRs and synaptic transmissions [18,46,61-66]. Interestingly, a gain-of-function mutation in the chrna1/AChR1 gene is reported to over-contract the muscle that induces mechani-cal damage of trunk muscles [67]. In rapsyn mutants, the postsynaptic AChR clusters are diffusely distributed and the mutant embryos are less active [46,64,68]. Similarly, zebrafish ennui mutants show reduced AChR aggregation at the NMJ and slow swimming in escape response, but the responsible gene for this mutation has not yet been determined [69]. In zebrafish, there are at least two functionally distinct splic-ing variants of MuSK [70]. A short form of MuSK regulates motor axon guidance [71,72], whereas a long form is involved in neuromuscular synapse formation in the focal regions of myotomes [73]. In zebrafish MuSK-null mutants, the focal synapses at equatorial locations are absent, whereas nonfocal synaptic contacts are formed in the vertical myoseptum. The AChR clustering at the nonfocal sites in fish depends on alpha-dystroglycan [73]. Although in choline acetyltransferase hypomorph, the postsynaptic AChRs are normally clustered, cholinergic transmissions at the NMJ are reduced and the escape response is attenuated [74]. In zebrafish acetylcholin-esterase (AChE) mutants, postsynaptic AChR clusters are initially formed, but are eventually reduced during develop-ment [75,76]. In accordance with the postsynaptic defects, the AChE mutants display progressive impairment of locomotion. These zebrafish mutants showing defects in the postsynaptic AChR clustering may be useful animal models of human NMJ disease.

    4. Chemicalbiologyinzebrafish

    Zebrafish is the only vertebrate available for drug screens on a laboratory scale.It has several advantages for discovering small molecules. First, zebrafish is prolific and thus suitable for high throughput assays. A pair of adult zebrafish can lay 100 200 fertilized eggs in a morning. As it is neither expensive nor difficult to maintain hundreds of adult fish in a laboratory, researchers can easily collect several embryos for screening. They are only 1 4 mm long during embryonic and early larval stages (up to 5 days postfertiliza-tion), when most of the internal organs are formed. Second, zebrafish live in water. In the embryonic and early larval stages, they do not eat food but receive nutrients from their yolk sac. Instead of preparing a drug-containing diet, researchers can dump drugs into the water. However, not all compounds may be permeable for embryos. Third, transgenic embryos, which express a certain marker gene, and mutant embryos are useful for chemical genetics. Taking advantage of transparency, a number of transgenic zebrafish have been generated that express GFP (green fluorescent protein), RFP (red fluorescent protein) or other fluorescent derivatives under control of tissue-specific, stress-inducible or cral 4-inducible promoters [77-81]. These transgenic embryos make it possible to visualize normal development of internal organs as well as the onset of physiological

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    changes under stress condition. The mutagenesis screens have so far identified > 4,000 mutants that show early developmental defects. In addition to the forward genetics, retrovirus- or transposon-mediated insertional mutagenesis and targeting induced local lesions in genome (TILLING) were established to generate zebrafish mutants [82-87]. More recently, gene targeting techniques using designed zinc-finger proteins have been developed [88,89]. These mutants may be useful for drug screening to find compounds that mitigate the mutant phenotypes. Fourth, antisense knockdown is applicable to any gene in the embryonic stage. Injection of antisense morpholino oligonucleotides into fertilized eggs can effectively block the translation or splicing of the target transcripts [90]. This loss of function assay may be useful to easily confirm the drug target [91].

    The first high throughput forward chemical screen using zebrafish embryos in 96-well plates has been done to find compounds that cause developmental malformations [92]. Three zebrafish embryos were placed in a well and small molecules were added to the arrayed embryos. Induced developmental defects were evaluated visually with a dissecting microscopy at 1, 2 and 3 days. Although, 2% of the small molecules were lethal, several novel compounds that affect development of the CNS, cardiovascular system, neural crest and ear were identified. Some other screens also identified compounds by focusing on protection from drug-induced cell death or developmental perturbations [93,94].

    Intriguingly, chemical screens to identify small molecules that can reverse developmental defects have also been done using zebrafish mutants. Zebrafish gridlock mutants have a hypomorphic mutation in hesr2 (hairy and enhancer of split related 2) gene that changes the stop codon to glycine and adds 44 amino acids at the C terminus [95]. The hesr2 prod-ucts function as a Notch effector in angioblast precursors and determine artery versus vein fate [96]. The gridlock mutants show malformation of dorsal aorta that results in interruption of blood flow to the trunk and tail [97]. Out of 5,000 drugs, 2 structurally related compounds were identified that rescue circulation to the tail. In another chemical suppres-sor screening, one novel compound, which canceled mitotic arrest in bmyb (myeloblastosis oncogene-like 2) mutants, was discovered in 16,000 small molecules [98]. These successful

    screenings show that zebrafish is useful in the discovery of novel drugs in vivo.

    5. Conclusion

    In the 1980s, zebrafish emerged in developmental study as a new animal model.As zebrafish is amenable to large-scale mutagenesis, many developmental biologists applied it to forward genetics. In the past decade, several useful genomic tools such as transgenics, gene inactivation and genome information have been developed that in turn expanded zebrafish research into many fields including neuroscience, immunology and cancer biology.Several zebrafish mutants that were relevant to many aspects of biological process have been generated. Some mutants were established as disease models with respect to the responsible gene and pathology. What can we do with these small patients? One of the most promising uses of zebrafish mutants may be high throughput screening of small molecules in vivo. Chemical screenings have successfully identified compounds that suppress mutant phenotypes. Zebrafish mutants generated either by forward mutagenesis or by reverse genetics such as TILLING or gene disruption by zinc-finger protein will be useful for drug discovery. If we find a new therapeutic molecule in zebrafish, its molecular target will be identified by in vitro analysis.

    6. Expertopinion

    Many zebrafish mutants that show neuromuscular and/or muscle defects have been identified by forward screening. Most of the mutants display muscle weakness or abnormal behavior that can be seen in human motor disorder. Although chemical genetics using these muscle mutants has not been reported, motility-based chemical screen will provide us novel therapeutic agents that mitigate human motor disorder such as muscle dystrophy and atrophy.


    The author states no conflict of interest and has received no payment in preparation of this manuscript.

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    ExpertOpin.DrugDiscov.(2009) 4(5) 511

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    AffiliationHiromi HirataAuthor for correspondenceNagoya University, Graduate School of Science, Proof to Hiromi Hirata Furo-cho, Chikusa-ku, Nagoya 464-8602, Japan Tel: +81 52 789 2980; Fax: +81 52 789 2979; E-mail: hhirata@bio.nagoya-u.ac.jp


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