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
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
2. Development of motility
3. Zebrafish motility mutants
caused by muscle or
neuromuscular junction defects
4. Chemical biology in zebrafish
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.
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 .
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 . These sensory neurons project to interneuronal networks located in the spinal cord and hindbrain to produce motor rhythm . 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 . 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 . Increased cytosolic Ca2+ activates troponin C that initiates actin/myosin sliding, thus causing muscle contraction . 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 . As activation of muscle is caused by rhythmic motor outputs, slow and fast bilateral alternation of muscle contractions generates coiling and swimming behavior, respectively.
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 . 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 . 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 . 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 . The mutant embryos lack both spontaneous and stimulus-evoked muscle contractility. Similarly, unc45b mutations impair skeletal myofibril formation and locomotion . 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 . 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 , 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 . 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 , 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 . 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 , 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 . 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 . 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 . In zebrafish, there are at least two functionally distinct splic-ing variants of MuSK . 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 . 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 . Although in choline acetyltransferase hypomorph, the postsynaptic AChRs are normally clustered, cholinergic transmissions at the NMJ are reduced and the escape response is attenuated . 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.
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 . This loss of function assay may be useful to easily confirm the drug target .
The first high throughput forward chemical screen using zebrafish embryos in 96-well plates has been done to find compounds that cause developmental malformations . 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 . The hesr2 prod-ucts function as a Notch effector in angioblast precursors and determine artery versus vein fate . The gridlock mutants show malformation of dorsal aorta that results in interruption of blood flow to the trunk and tail . 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 . These successful
screenings show that zebrafish is useful in the discovery of novel drugs in vivo.
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.
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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Bibliography1. Pourqui O. Vertebrate somitogenesis.
Annu Rev Cell Dev Biol 2001;17:311-50
2. Saga Y, Takeda H. The making of the somite: molecular events in vertebrate segmentation. Nat Rev Genet 2001;2:835-45
3. Currie PD, Ingham PW. Induction of a specific muscle cell type by a hedgehog-like protein in zebrafish. Nature 1996;382:452-5
4. Blagden CS, Currie PD, Ingham PW, Hughes SM. Notochord induction of zebrafish slow muscle mediated by sonic hedgehog. Genes Dev 1997;11:2163-75
5. Devoto SH, Melanon E, Eisen JS, Westerfield M. Identification of separate slow and fast muscle precursor cells in vivo, prior to somite formation. Development 1996;122:3371-80
6. Stellabotte F, Dobbs-McAuliffe B, Fernndez DA, et al. Dynamic somite cell rearrangements lead to distinct waves of myotome growth. Development 2007;134:1253-7
7. Ochi H, Westerfield M. Signaling networks that regulate muscle development: lessons from zebrafish. Dev Growth Differ 2007;49:1-11
8. Nguyen PV, Aniksztejn L, Catarsi S, Drapeau P. Maturation of neuromuscular transmission during early development in zebrafish. J Neurophysiol 1999;81:2852-61
9. Buss RR, Drapeau P. Synaptic drive to motoneurons during fictive swimming in the developing zebrafish. J Neurophysiol 2001;86:197-210
10. Luna VM, Brehm P. An electrically coupled network of skeletal muscle in zebrafish distributes synaptic current. J Gen Physiol 2006;128:89-102
11. Saint-Amant L, Drapeau P. Time course of the development of motor behaviors in the zebrafish embryo. J Neurobiol 1998;37:622-32
12. Downes GB, Granato M. Supraspinal input is dispensable to generate glycine-mediated locomotive behaviors in the zebrafish embryo. J Neurobiol 2006;66:437-51
13. Metcalfe WK, Myers PZ, Trevarrow B, et al. Primary neurons that express the L2/HNK-1 carbohydrate during early development in the zebrafish. Development 1990;110:491-504
14. Drapeau P, Saint-Amant L, Buss RR, et al. Development of the locomotor network in zebrafish. Prog Neurobiol 2002;68:85-111
15. Svoboda KR, Linares AE, Ribera AB. Activity regulates programmed cell death of zebrafish Rohon-Beard neurons. Development 2001;128:3511-20
16. Fetcho JR, Higashijima S, McLean DL. Zebrafish and motor control over the last decade. Brain Res Rev 2008;57:86-93
17. Buss RR, Drapeau P. Activation of embryonic red and white muscle fibers during fictive swimming in the developing zebrafish. J Neurophysiol 2002;87:1244-51
18. Li W, Ono F, Brehm P. Optical measurements of presynaptic release in mutant zebrafish lacking postsynaptic receptors. J Neurosci 2003;23:10467-74
19. Catterall WA. Structure and regulation of voltage-gated Ca2+ channels. Annu Rev Cell Dev Biol 2000;16:521-55
20. Franzini-Armstrong C, Jorgensen AO. Structure and development of E-C coupling units in skeletal muscle. Annu Rev Physiol 1994;56:509-34
21. Gordon AM, Homsher E, Regnier M. Regulation of contraction in striated muscle. Physiol Rev 2000;80:853-924
22. MacLennan DH. Ca2+ signaling and muscle disease. Eur J Biochem 2000;267:5291-7
23. Streisinger G, Walker C, Dower N, et al. Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature 1981;291:293-6
24. Chakrabarti S, Streisinger G, Singer F, Walker C. Frequency of gamma-ray induced specific locus and recessive lethal mutations in mature germ cells of the zebrafish, BRACHYDANIO RERIO. Genetics 1983;103:109-23
25. Walker C, Streisinger G. Induction of mutations by gamma-rays in pregonial germ cells of zebrafish embryos. Genetics 1983;103:125-36
26. Grunwald DJ, Streisinger G. Induction of recessive lethal and specific locus mutations in the zebrafish with ethyl nitrosourea. Genet Res 1992;59:103-16
27. Grunwald DJ, Eisen JS. Headwaters of the zebrafishemergence of a new model vertebrate. Nat Rev Genet 2002;3:717-24
28. Haffter P, Granato M, Brand M, et al. The identification of genes with unique
and essential functions in the development of the zebrafish, Danio rerio. Development 1996;123:1-36
29. Driever W, Solnica-Krezel L, Schier AF, et al. A genetic screen for mutations affecting embryogenesis in zebrafish. Development 1996;123:37-46
30. Granato M, van Eeden FJ, Schach U, et al. Genes controlling and mediating locomotion behavior of the zebrafish embryo and larva. Development 1996;123:399-413
31. Bassett DI, Bryson-Richardson RJ, Daggett DF, et al. Dystrophin is required for the formation of stable muscle attachments in the zebrafish embryo. Development 2003;130:5851-60
32. Guyon JR, Goswami J, Jun SJ, et al. Genetic isolation and characterization of a splicing mutant of zebrafish dystrophin. Hum Mol Genet 2009;18:202-11
33. Hall TE, Bryson-Richardson RJ, Berger S, et al. The zebrafish candyfloss mutant implicates extracellular matrix adhesion failure in laminin 2-deficient congenital muscular dystrophy. Proc Natl Acad Sci USA 2007;104:7092-7
34. Xu X, Meiler SE, Zhong TP, et al. Cardiomyopathy in zebrafish due to mutation in an alternatively spliced exon of titin. Nat Genet 2002;30:205-9
35. Steffen LS, Guyon JR, Vogel ED, et al. The zebrafish runzel muscular dystrophy is linked to the titin gene. Dev Biol 2007;309:180-92
36. Bassett DI, Currie PD. The zebrafish as a model for muscular dystrophy and congenital myopathy. Hum Mol Genet 2003;15:R265-270
37. Kunkel LM, Bachrach E, Bennett RR, et al. Diagnosis and cell-based therapy for Duchenne muscular dystrophy in humans, mice, and zebrafish. J Hum Genet 2006;51:397-406
38. Steffen LS, Guyon JR, Vogel ED, et al. Zebrafish orthologs of human muscular dystrophy genes. BMC Genomics 2007;8:79
39. Hawkins TA, Haramis AP, Etard C, et al. The ATPase-dependent chaperoning activity of Hsp90a regulates thick filament formation and integration during skeletal muscle myofibrillogenesis. Development 2008;135:1147-56
40. Etard C, Behra M, Fischer N, et al. The UCS factor Steif/Unc-45b interacts
512 ExpertOpin.DrugDiscov.(2009) 4(5)
with the heat shock protein Hsp90a during myofibrillogenesis. Dev Biol 2007;308:133-43
41. Wohlgemuth SL, Crawford BD, Pilgrim DB. The myosin co-chaperone UNC-45 is required for skeletal and cardiac muscle function in zebrafish. Dev Biol 2007;303:483-92
42. Du SJ, Li H, Bian Y, Zhong Y. Heat-shock protein 901 is required for organized myofibril assembly in skeletal muscles of zebrafish embryos. Proc Natl Acad Sci USA 2008;105:554-9
43. Etard C, Roostalu U, Strhle U. Shuttling of the chaperones Unc45b and Hsp90a between the A band and the Z line of the myofibril. J Cell Biol 2008;180:1163-75
44. Felsenfeld AL, Walker C, Westerfield M, et al. Mutations affecting skeletal muscle myofibril structure in the zebrafish. Development 1990;108:443-59
45. Felsenfeld AL, Curry M, Kimmel CB. The fub-1 mutation blocks initial myofibril formation in zebrafish muscle pioneer cells. Dev Biol 1991;148:23-30
46. Ono F, Higashijima S, Shcherbatko A, et al. Paralytic zebrafish lacking acetylcholine receptors fail to localize rapsyn clusters to the synapse. J Neurosci 2001;21:5439-48
47. Schredelseker J, Di Biase V, Obermair GJ, et al. The b 1a subunit is essential for the assembly of dihydropyridine-receptor arrays in skeletal muscle. Proc Natl Acad Sci USA 2005;102:17219-24
48. Zhou W, Saint-Amant L, Hirata H, et al. Non-sense mutations in the dihydropyridine receptor b1 gene, CACNB1, paralyze zebrafish relaxed mutants. Cell Calcium 2006;39:227-36
49. Schredelseker J, Dayal A, Schwerte T, et al. Proper restoration of excitation-contraction coupling in the dihydropyridine receptor b 1-null zebrafish relaxed is an exclusive function of the b 1a subunit. J Biol Chem 2008;284:1242-51
50. Ptcek LJ, Tawil R, Griggs RC, et al. Dihydropyridine receptor mutations cause hypokalemic periodic paralysis. Cell 1994;77:863-8
51. Hirata H, Watanabe T, Hatakeyama J, et al. Zebrafish relatively relaxed mutants have a ryanodine receptor defect, show slow swimming and provide a model of
multi-minicore disease. Development 2007;134:2771-81
52. Zhang Y, Chen HS, Khanna VK, et al. A mutation in the human ryanodine receptor gene associated with central core disease. Nat Genet 1993;5:46-50
53. Jungbluth H, Beggs A, Bnnemann C, et al. 111th ENMC international workshop on multi-minicore disease. 2nd international MmD workshop, 9-11 November 2002, Naarden, The Netherlands. Neuromuscul Disord 2004;14:754-66
54. Monnier N, Ferreiro A, Marty I, et al. A homozygous splicing mutation causing a depletion of skeletal muscle RYR1 is associated with multi-minicore disease congenital myopathy with ophthalmoplegia. Hum Mol Genet 2003;12:1171-8
55. Hirata H, Saint-Amant L, Waterbury J, et al. accordion, a zebrafish behavioral mutant, has a muscle relaxation defect due to a mutation in the ATPase Ca2+ pump SERCA1. Development 2004;131:5457-68
56. Gleason MR, Armisen R, Verdecia MA, et al. A mutation in serca underlies motility dysfunction in accordion zebrafish. Dev Biol 2004;276:441-51
57. Odermatt A, Taschner PE, Khanna VK, et al. Mutations in the gene-encoding SERCA1, the fast-twitch skeletal muscle sarcoplasmic reticulum Ca2+ ATPase, are associated with Brody disease. Nat Genet 1996;14:191-4
58. Sanes JR, Lichtman JW. Induction, assembly, maturation and maintenance of a postsynaptic apparatus. Nat Rev Neurosci 2001;2:791-805
59. Kim N, Stiegler AL, Cameron TO, et al. Lrp4 is a receptor for Agrin and forms a complex with MuSK. Cell 2008;135:334-42
60. Zhang B, Luo S, Wang Q, et al. LRP4 serves as a coreceptor of agrin. Neuron 2008;60:285-97
61. Westerfield M, Liu DW, Kimmel CB, Walker C. Pathfinding and synapse formation in a zebrafish mutant lacking functional acetylcholine receptors. Neuron 1990;4:867-74
62. Sepich DS, Ho RK, Westerfield M. Autonomous expression of the nic1 acetylcholine receptor mutation in zebrafish muscle cells. Dev Biol 1994;161:84-90
63. Sepich DS, Wegner J, OShea S, Westerfield M. An altered intron inhibits synthesis of the acetylcholine receptor -subunit in the paralyzed zebrafish mutant nic1. Genetics 1998;148:361-72
64. Ono F, Mandel G, Brehm P. Acetylcholine receptors direct rapsyn clusters to the neuromuscular synapse in zebrafish. J Neurosci 2004;24:5475-81
65. Panzer JA, Song Y, Balice-Gordon RJ. In vivo imaging of preferential motor axon outgrowth to and synaptogenesis at prepatterned acetylcholine receptor clusters in embryonic zebrafish skeletal muscle. J Neurosci 2006;26:934-47
66. Epley KE, Urban JM, Ikenaga T, Ono F. A modified acetylcholine receptor d-subunit enables a null mutant to survive beyond sexual maturation. J Neurosci 2008;28:13223-31
67. Lefebvre JL, Ono F, Puglielli C, et al. Increased neuromuscular activity causes axonal defects and muscular degeneration. Development 2004;131:2605-18
68. Ono F, Shcherbatko A, Higashijima S, et al. The zebrafish motility mutant twitch once reveals new roles for rapsyn in synaptic function. J Neurosci 2002;22:6491-8
69. Saint-Amant L, Sprague SM, Hirata H, et al. The zebrafish ennui behavioral mutation disrupts acetylcholine receptor localization and motor axon stability. Dev Neurobiol 2008;68:45-61
70. Zhang J, Lefebvre JL, Zhao S, Granato M. Zebrafish unplugged reveals a role for muscle-specific kinase homologs in axonal pathway choice. Nat Neurosci 2004;7:1303-9
71. Zhang J, Granato M. The zebrafish unplugged gene controls motor axon pathway selection. Development 2000;127:2099-111
72. Zhang J, Malayaman S, Davis C, Granato M. A dual role for the zebrafish unplugged gene in motor axon pathfinding and pharyngeal development. Dev Biol 2001;240:560-73
73. Lefebvre JL, Jing L, Becaficco S, et al. Differential requirement for MuSK and dystroglycan in generating patterns of neuromuscular innervation. Proc Natl Acad Sci USA 2007;104:2483-8
74. Wang M, Wen H, Brehm P. Function of neuromuscular synapses in the zebrafish
ExpertOpin.DrugDiscov.(2009) 4(5) 513
choline-acetyltransferase mutant bajan. J Neurophysiol 2008;100:1995-2004
75. Behra M, Cousin X, Bertrand C, et al. Acetylcholinesterase is required for neuronal and muscular development in the zebrafish embryo. Nat Neurosci 2002;5:111-8
76. Downes GB, Granato M. Acetylcholinesterase function is dispensable for sensory neurite growth but is critical for neuromuscular synapse stability. Dev Biol 2004;270:232-45
77. Amsterdam A, Lin S, Hopkins N. The aequorea victoria green fluorescent protein can be used as a reporter in live zebrafish embryos. Dev Biol 1995;171:123-9
78. Higashijima S, Okamoto H, Ueno N, et al. High-frequency generation of transgenic zebrafish which reliably express GFP in whole muscles or the whole body by using promoters of zebrafish origin. Dev Biol 1997;192:289-99
79. Long Q, Meng A, Wang H, et al. GATA-1 expression pattern can be recapitulated in living transgenic zebrafish using GFP reporter gene. Development 1997;124:4105-11
80. Scheer N, Campos-Ortega JA. Use of the Gal4-UAS technique for targeted gene expression in the zebrafish. Mech Dev 1999;80:153-8
81. Halloran MC, Sato-Maeda M, Warren JT, et al. Laser-induced gene expression in specific cells of transgenic zebrafish. Development 2000;127:1953-60
82. Lin S, Gaiano N, Culp P, et al. Integration and germ-line transmission of a
pseudotyped retroviral vector in zebrafish. Science 1994;265:666-9
83. Gaiano N, Amsterdam A, Kawakami K, et al. Insertional mutagenesis and rapid cloning of essential genes in zebrafish. Nature 1996;383:829-32
84. Kawakami K, Shima A, Kawakami N. Identification of a functional transposase of the Tol2 element, an Ac-like element from the Japanese medaka fish, and its transposition in the zebrafish germ lineage. Proc Natl Acad Sci USA 2000;97:11403-8
85. Kawakami K, Takeda H, Kawakami N, et al. A transposon-mediated gene trap approach identifies developmentally regulated genes in zebrafish. Dev Cell 2004;7:133-44
86. Sivasubbu S, Balciunas D, Davidson AE, et al. Gene-breaking transposon mutagenesis reveals an essential role for histone H2afza in zebrafish larval development. Mech Dev 2006;123:513-29
87. Wienholds E, Schulte-Merker S, Walderich B, Plasterk RH. Target-selected inactivation of the zebrafish rag1 gene. Science 2002;297:99-102
88. Meng X, Noyes MB, Zhu LJ, et al. Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Nat Biotechnol 2008;26:695-701
89. Doyon Y, McCammon JM, Miller JC, et al. Heritable targeted gene disruption in zebrafish using designed zinc-finger nucleases. Nat Biotechnol 2008;26:702-8
90. Nasevicius A, Ekker SC. Effective targeted gene knockdown in zebrafish. Nat Genet 2000;26:216-20
91. Zon LI, Peterson RT. In vivo drug discovery in the zebrafish. Nat Rev Drug Discov 2005;4:35-44
92. Peterson RT, Link BA, Dowling JE, Schreiber SL. Small molecule developmental screens reveal the logic and timing of vertebrate development. Proc Natl Acad Sci USA 2000;97:12965-9
93. Owens KN, Santos F, Roberts B, et al. Identification of genetic and chemical modulators of zebrafish mechanosensory hair cell death. PLoS Genet 2008;4:e1000020
94. Sachidanandan C, Yeh JR, Peterson QP, Peterson RT. Identification of a novel retinoid by small molecule screening with zebrafish embryos. PLoS ONE 2008;3:e1947
95. Zhong TP, Rosenberg M, Mohideen MA, et al. gridlock, an HLH gene required for assembly of the aorta in zebrafish. Science 2000;287:1820-4
96. Zhong TP, Childs S, Leu JP, Fishman MC. Gridlock signalling pathway fashions the first embryonic artery. Nature 2001;414:216-20
97. Peterson RT, Shaw SY, Peterson TA, et al. Chemical suppression of a genetic mutation in a zebrafish model of aortic coarctation. Nat Biotechnol 2004;22:595-9
98. Stern HM, Murphey RD, Shepard JL, et al. Small molecules that delay S phase suppress a zebrafish bmyb mutant. Nat Chem Biol 2005;1:366-70
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: firstname.lastname@example.org