1. Department of Microbiology, Immunology and Molecular Genetics,
2. Department of Molecular, Cell, and Developmental Biology
3. Molecular Biology Institute, University of California, Los Angeles, California 90095, USA
4. Biological Imaging Center, Beckman Institute, California Institute of Technology, Pasadena, California 91125, USA
5. These authors contributed equally to this work.

Correspondence to: Kent L. Hill^1,3 Correspondence and requests for materials should be addressed to K.L.H. (Email: kenthill@mednet.ucla.edu).

Cilia are evolutionarily conserved organelles that perform motility, sensory and transport functions and are required for normal vertebrate development and physiology^{2, 3, 4, 5}. As such, cilium defects underlie a broad spectrum of human diseases^{4, 5}. Among the roles of ciliated organs in vertebrate embryogenesis, the contribution of cilia to inner-ear development remains poorly understood. In the zebrafish, Danio rerio, it has been proposed that beating cilia participate in the biogenesis of otoliths⁶, which are analogous to otoconia in the otolithic membrane of human ears. These biomineralized particles provide an inertial mass that facilitates deflection of underlying microvilli and cilia, thereby initiating signalling events that allow the brain to detect sound, gravity and linear acceleration^{1, 7, 8, 9}. During early development, nascent otoliths are formed from a pool of precursor particles and tethered to cilia in the otic vesicle⁶. So far, direct evidence for the necessity of ciliary motility in this process is lacking.

In protists, ciliary motility is controlled by the Dynein Regulatory Complex (DRC), which regulates axonemal dynein activity in response to signals from the radial spokes and central pair apparatus^{10, 11, 12, 13, 14, 15}. The DRC subunit trypanin is conserved across diverse phyla^{15, 16, 17, 18} and the vertebrate (human) trypanin homologue, growth arrest-specific 8 (here called GAS8, in line with the HUGO database, but also, and originally, designated GAS11) is a microtubule-binding protein localized to regions of dynein regulation in mammalian cells^{19, 20, 21}. So far, however, a requirement for GAS8 and the DRC in vertebrates has not been established. We identified a single trypanin homologue in zebrafish encoding a protein that is 63.8% identical to the human GAS8 protein and 32.0% identical to trypanin from Trypanosoma brucei (Fig. 1). The sequence identity and conserved genomic structure (Fig. 2a)^{15, 22} indicate that this zebrafish protein, designated Gas8, is indeed a member of this conserved family of dynein regulatory proteins^{13, 15}. Maternal gas8 transcripts are ubiquitous throughout the embryo during early development (Fig. 1b, c). By the 12-somite stage, however, expression becomes concentrated in the developing ears (Fig. 1d, arrow) and this persists through the 18- to 20-somite stage (Fig. 1e, Supplementary Fig. 1). Transcripts are also clearly present in the brain, neural tube and pronephric ducts (Fig. 1f–h). Therefore, gas8 is expressed in ciliated tissues during zebrafish organogenesis.

Figure 1: gas8 is expressed in ciliated tissues.

a, Protein sequence alignment of trypanin/GAS8 from Homo sapiens, Danio rerio and Trypanosoma brucei. Yellow highlighting indicates absolutely conserved residues, blue indicates residues conserved in two homologues and green indicates residues that are conservative substitutions. The boxed region indicates a conserved RNYFQERDK stretch that is found in every known trypanin/GAS8 homologue¹⁷. The conserved microtubule binding domain 'GMAD' and the regulatory domain 'IMAD'¹⁹ are indicated with blue and black overlines, respectively. b–h, In situ hybridizations show the gas8 expression pattern during the first 24 h of embryonic development. Developing ears (black arrows), neural tube (open arrowhead) and pronephric ducts (filled arrowheads) are shown. Developmental stages are indicated in each panel; S, somite. h is an enlargement of the boxed region in g.

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Figure 2: gas8 morphants exhibit developmental defects.

a, Intron/exon structure of the gas8 locus, which encodes a predicted 1.54-kb transcript. The positions of splice blocking (SP MO) and translation blocking (AUG MO) morpholino oligonucleotides are shown. b, RNA from wild-type (WT) and gas8 splice morphant embryos was analysed by PCR with reverse transcription (RT–PCR) using a forward primer (a) in the second exon and a reverse primer in either the second intron (b) or the third exon (c). In the morphant, blocking of the exon-2 splice donor site leads to a 315-bp RT–PCR product with the first primer set and no product with the second primer set. Controls for RT–PCR were provided by amplification of a 95-bp fragment of gapdh. c–f, gas8 morphants have a variety of defects: overall morphology of controls (c) and gas8 morphants (d) at 24 h.p.f.; detail of control (e) and morphant (f) embryos at 3 days post-fertilization (d.p.f.) showing hydrocephaly (white arrow), pericardial oedema (open arrowhead), disorganized somites and otolith abnormalities (black arrow). g, Quantitative analysis of otolith defects at 3 d.p.f. The relative number of fish having the indicated defect is shown for uninjected embryos (stippled bars; n = 324, five experiments), embryos injected with control MO (white bars; n = 167, two experiments), SP MO (grey bars; n = 89, two experiments), AUG MO (black bars; n = 96, two experiments) or co-injected with AUG MO and 250 pg in-vitro-transcribed gas8 mRNA (hatched bars; n = 225, two experiments). Error bars, s.d. h–p, Panels show the spectrum of otolith defects observed in gas8 morphants at 3 d.p.f. (h–m) and earlier times (n–p): normal otoliths (h); a single otolith (i); ectopic, fused and small otoliths (j–m); and nascent otoliths in control (n, 27 h.p.f.) and gas8 morphant (o, 24 h.p.f.; p, 27 h.p.f.) embryos. Scale bars, 30 m (h–m); 20 m (n–p). White arrowhead indicates ectopic and fused otoliths in the gas8 morphant. Embryos were injected with 6 ng (AUG MO), 4 or 5 ng (SP MO), 6 ng (standard control MO) or 6 ng (mismatch AUG MO) morpholinos.

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To determine how loss of gas8 expression affects zebrafish development we employed antisense morpholino oligonucleotides (Fig. 2). Hydrocephaly, neural tube cell death and left–right axis defects are common in ciliary morphants and mutants³ and were evident in gas8 morphants (Fig. 2, Supplementary Fig. 2). Given the high expression of gas8 in the otic vesicle, we examined ear development more closely. By two days post-fertilization, inner-ear atrophy was evident gas8 morphants. The length along the antero-posterior axis was 30% less than in control embryos (morphants, 52  6 m, n = 15; control, 73  7 m, n = 8; 24–27 hours post-fertilization (h.p.f.); see, for example, Figs 2e, f, 3a, b). By three days post-fertilization, exactly two otoliths were present in control zebrafish, one at the anterior end and one at the posterior end of the otic vesicle, positioned ventrally to a semicircular canal⁶. By contrast, gas8 morphants had abnormal numbers of otoliths, fused otoliths, abnormally positioned otoliths and small otoliths (Fig. 2g–m). Examination of 24-h.p.f. and 27-h.p.f. embryos showed that the same spectrum of defects is evident during nascent otolith formation (Fig. 2n–p). The otolith phenotype is 95–100% penetrant and co-injection of in-vitro-transcribed gas8 messenger RNA carrying five base-pair mismatch mutations to prevent morpholino hybridization rescued the defect in a majority of embryos (Fig. 2g). Because inner ear patterning is tightly linked with neuraxis patterning²³, we analysed markers for the hindbrain (egr2b), midbrain (eng2a), forebrain (otx2), inner-ear anterio-ventral area (fgf8a)²⁴ and inner-ear anterior/posterior extremity (bmp4)²⁵. We did not detect differences between controls and gas8 morphants (data not shown). Therefore, otolith defects are not due to abnormal neuraxis or inner-ear patterning.

Figure 3: Gas8 is required for tether cilia motility.

a, b, Cilia in control (a) and gas8 morphant (b) embryos at 24 h.p.f., visualized by immunofluorescence labelling with anti-acetylated tubulin antibodies (Ac. Tub.). Arrowheads indicate the location of the tether cilia and arrows indicate short cilia. Scale bars, 10 m. c–n, Tether cilia are motile in control embryos, but not in gas8 morphants. Bright-field snapshot images from high-speed videos of cilia in controls (c, d) and gas8 morphants (e, f), showing two steps of the cilia beat cycle with a 15-ms interval (half the period of a beat). g–j, Time-to-colour merge of six frames encompassing 15 ms of the cilia motion immediately preceding the still images shown in c–f, respectively. Cilia position in time is marked by different colours following the colour bar. When merged, moving objects are visible in the corresponding colours, whereas immotile objects only show background and noise. k–n, Diagrams showing cilia and otolith motion in control (k, l) and immotility in gas8 morphant (m, n) embryos with three time points along half the period (15 ms) of the cilia beat cycle (see colour bar). Still images from control (c, d) and gas8 morphant (e, f) embryos are taken from Supplementary Movies 3 and 5, respectively. Scale bars, 1 m (c–f). Arrowheads point to tether cilia.

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We next asked whether otolith defects were due to improper formation or placement of cilia in the otic vesicle. By 24 h.p.f., two classes of cilia were visible in the otic vesicle⁶ (Fig. 3a, Supplementary Movie 1). Numerous short cilia were dispersed throughout the otic vesicle, and small patches of longer, 'tether' cilia were found exclusively at the anterior and posterior poles. Between 19 and 27 h.p.f., small precursor particles coalesced on tether cilia to form anterior and posterior otoliths⁶. We examined cilia distribution and size in morphant embryos during the critical developmental window of 19–27 h.p.f., when cilia are postulated to function in otolith assembly. At the 20-somite stage (corresponding to 19 h.p.f. in wild type), control and morphant embryos had cilia in the developing otic vesicle (data not shown). gas8 morphants were slightly developmentally delayed, a common feature of morphant fish. Hence, we staged embryos based on developmental progression defined in ref. 26. By 24 h.p.f., both short cilia and tether cilia were distinguishable at the correct locations in control and gas8 morphant embryos (Fig. 3b, Supplementary Movie 2). Otic vesicle size was reduced as noted above, but we did not detect major length differences of cilia between control (tether, 5.9  0.2 m; short, 1.4  0.1 m; n = 5 embryos) and morphant (tether, 5.9  0.4 m; short, 1.2  0.2 m; n = 7 embryos) embryos. At later stages, tether cilia persisted whereas short cilia began to disappear as expected⁶ in both control and morphant embryos (data not shown). Cilia were also observed in the pronephric ducts and neural tube in gas8 morphants (data not shown). Therefore, loss of gas8 expression did not prevent formation, maintenance or correct positioning of cilia.

Because protist GAS8 homologues, trypanin in T. brucei and paralyzed flagella 2 in Chlamydomonas reinhardtii, are specifically involved in controlling ciliary beat^{13, 14, 15}, we examined cilia motility directly using in vivo, high-speed video microscopy. In all control embryos, one to three beating tether cilia were detected near each nascent otolith (Fig. 3, Supplementary Fig. 3, Supplementary Movies 3–5). Beating cilia directly bore the forming otolith or were located nearby (5–10 m distant), often causing the otolith itself to oscillate (Supplementary Movies 3, 5). These cilia beat with a frequency of 34  6 Hz (n = 20) at 24 h.p.f. Short cilia were not motile (Supplementary Movie 6). This contrasts with a previous report suggesting that tether cilia are immotile whereas short cilia distributed throughout the ear are motile⁶. The reason for this discrepancy is not clear, but it is probably due to technical challenges associated with imaging cilia motility, which was inferred indirectly in the earlier work and imaged directly here. In gas8 morphants at every stage examined (19–27 h.p.f.), a majority of embryos displayed immotile tether cilia (60%, n = 30; Fig. 3, Supplementary Fig. 3, Supplementary Movies 7–8). Commonly, gas8 morphants displayed ectopic otoliths located at the base of non-motile tether cilia (Fig. 2o). In some cases, morphants harboured ectopic beating cilia (23%, n = 30). To confirm that the ear phenotype was due to cilia immotility and not loss of another, unknown gas8 function, we knocked down two genes directly involved in cilia motility: the gene for leucine-rich repeat-containing 50 protein, lrrc50, an outer-arm dynein subunit shown previously to be required for cilia motility^{27, 28}; and the left–right dynein gene dnah9, a well-characterized motor protein involved in cilia motility²⁹. The same results were obtained upon knockdown of lrrc50 (Supplementary Figs 4, 5). Furthermore, simultaneous knockdown of lrrc50 and dnah9 had a synergistic effect, causing more significant motility and otolith defects than either single knockdown alone. These treatments neither affected brain development nor triggered neural tube cell death and pericardial oedema. In all cases, abnormal otolith size, number and positioning were directly correlated with defective ciliary motility in morphants (Supplementary Fig. 4).

Otoliths are composite crystals assembled from a common pool of small, precursor particles. We hypothesized that otolith defects in cilia morphants arise as a consequence of abnormal fluid flow and the concomitant failure to properly direct precursor particle movements. To test this hypothesis, we examined fluid flow patterns in the otic vesicle by tracking otolith precursors at high temporal resolution. A direct correlation between cilia beat and fluid flow was observed (Fig. 4, Supplementary Movies 9–11). In control embryos, the motion of precursor particles near the otolith was non-random, whereas those further away from the otolith exhibited Brownian motion (Fig. 4g, h, Supplementary Movies 9–10), consistent with a previous report⁶. Cilia beating triggers a local flow in the vicinity of the otolith, attracting precursors at the base of tether cilia and propelling them towards the otolith (Supplementary Fig. 6, Supplementary Movies 9–10). By contrast, in gas8 morphants, particles near tether cilia exhibited purely diffusive behaviour (Fig. 4, Supplementary Movie 11). Therefore, the absence of ciliary beating limits particles to random motion, leading to formation of ectopic aggregates.

Figure 4: Tether cilia motility drives otolith biogenesis.

a–h, Particle tracking analysis demonstrates that cilium-dependent fluid dynamics drive precursor particle movement near the otolith. In control embryos, particle tracks (a) and displacements (b) show that particles near the otolith move by non-Brownian motion. Each track has a different colour. d, e, In gas8 morphants, particle tracks (d) and displacements (e) show decreased particle displacements in comparison with control. c, f, Mean particle displacement, r(t), plotted as a function of time. In control embryos, mean particle displacement is large and non-random, indicating diffusive transport. In gas8 morphants, mean particle displacement is small and random, indicating Brownian motion only. Error bars indicate the variance in the calculation of r(t). g, h, Particle tracking in control embryos shows that particle displacement is directly correlated with position relative to the otolith. g, particle tracks. h, Displacements of particles in regions I, II and III of g were calculated as a function of time and the average of the mean displacement, r(t), for each particle is shown. Net particle displacement decreases with increasing distance from tether cilia, indicating the reduction of the influence of ciliary beating. i–k, Diagrams depicting cilium-dependent otolith biogenesis. i, Tether cilia motility creates vortices that attract precursor particles. j, Cilia motility further serves to disperse particles locally and causes oscillation of the otolith, together facilitating uniform otolith growth. k, In gas8 morphants, absence of ciliary motility limits particles to Brownian motion. l, In wild-type embryos, the net consequence of tether cilia motility is that precursor particles are concentrated near the tethers, preferentially seeding otoliths at two poles of the otic vesicle. m, In gas8 morphants, loss of normal ciliary motility leads to formation of ectopic aggregates, non-uniform otolith growth and small particles that fail to coalesce into full-sized otoliths. Scale bars, 5 m.

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Our results demonstrate that Gas8 is required for normal motility of cilia in the otic vesicle and that ciliary motility is essential for normal ear development. The otic vesicle is a closed epithelial organ and fluid flow within this vesicle has been suggested to contribute to otolith formation⁶. Our study provides direct experimental evidence in support of this hypothesis. On the basis of high-speed video microscopy of cilia motility and quantitative analysis of precursor particle movements in wild-type and gas8 morphant embryos, we propose a new, cilium-dependent hydrodynamic mechanism for otolith biogenesis (Fig. 4). In this model, motility of tether cilia at the poles of the otic vesicle establishes a vortex that attracts otolith precursors (Fig. 4i, l), thereby biasing an otherwise random distribution of precursor particles and concentrating them near the two patches of tether cilia. This ensures preferential otolith seeding at the poles of the otic vesicle. At the otic vesicle poles, tether cilia motility further serves to disperse precursor particles locally and oscillation of the otolith increases effective contact area with precursors (Fig. 4j). Together, this prevents particles from sedimenting to form ectopic aggregates and promotes efficient uniform otolith growth. This model explains the different features of the otolith phenotype observed in gas8 morphants (Fig. 4k, m).

Our findings add to a growing list of developmental processes requiring fluid dynamic inputs for proper growth and patterning, further showing that epigenetic cues are part of the embryonic developmental program. In humans, hearing and balance defects are common among the elderly and are the most frequent sensory hereditary defects in newborns³⁰. Although human patients with ciliopathies have not generally been observed to have obvious hearing loss⁴, our results should stimulate investigations to look for more subtle inner-ear changes. To conclude, our studies demonstrate a requirement for motile cilia in vertebrate ear development and suggest that DRC subunits should be considered as candidates for disease genes contributing to ciliopathies in humans.

References

1. Sollner, C. & Nicolson, T. in Biomineralization: From Biology to Biotechnology and Medical Application 2nd edn (ed. Bauerlein, E.) 229–242 (Wiley-VCH, 2004)
2. Satir, P. & Christensen, S. T. Overview of structure and function of mammalian cilia. Annu. Rev. Physiol. 69, 377–400 (2007) | Article | PubMed | ISI | ChemPort |
3. Bisgrove, B. W. & Yost, H. J. The roles of cilia in developmental disorders and disease. Development 133, 4131–4143 (2006) | Article | PubMed | ChemPort |
4. Badano, J. L., Mitsuma, N., Beales, P. L. & Katsanis, N. The ciliopathies: an emerging class of human genetic disorders. Annu. Rev. Genomics Hum. Genet. 7, 125–148 (2006) | Article | PubMed | ISI | ChemPort |
5. Fliegauf, M., Benzing, T. & Omran, H. When cilia go bad: cilia defects and ciliopathies. Nature Rev. Mol. Cell Biol. 8, 880–893 (2007) | Article |
6. Riley, B. B., Zhu, C., Janetopoulos, C. & Aufderheide, K. J. A critical period of ear development controlled by distinct populations of ciliated cells in the zebrafish. Dev. Biol. 191, 191–201 (1997) | Article | PubMed | ISI | ChemPort |
7. Haddon, C. & Lewis, J. Early ear development in the embryo of the zebrafish, Danio rerio. J. Comp. Neurol. 365, 113–128 (1996) | Article | PubMed | ChemPort |
8. Hughes, I., Thalmann, I., Thalmann, R. & Ornitz, D. M. Mixing model systems: using zebrafish and mouse inner ear mutants and other organ systems to unravel the mystery of otoconial development. Brain Res. 1091, 58–74 (2006) | Article | PubMed | ChemPort |
9. Nicolson, T. The genetics of hearing and balance in zebrafish. Annu. Rev. Genet. 39, 9–22 (2005) | Article | PubMed | ChemPort |
10. Huang, B., Ramanis, Z. & Luck, D. J. Suppressor mutations in Chlamydomonas reveal a regulatory mechanism for Flagellar function. Cell 28, 115–124 (1982) | Article | PubMed | ISI | ChemPort |
11. Piperno, G., Mead, K. & Shestak, W. The inner dynein arms I2 interact with a "dynein regulatory complex" in Chlamydomonas flagella. J. Cell Biol. 118, 1455–1463 (1992) | Article | PubMed | ChemPort |
12. Gardner, L. C., O'Toole, E., Perrone, C. A., Giddings, T. & Porter, M. E. Components of a "dynein regulatory complex" are located at the junction between the radial spokes and the dynein arms in Chlamydomonas flagella. J. Cell Biol. 127, 1311–1325 (1994) | Article | PubMed | ChemPort |
13. Ralston, K. S., Lerner, A. G., Diener, D. R. & Hill, K. L. Flagellar motility contributes to cytokinesis in Trypanosoma brucei and is modulated by an evolutionarily conserved dynein regulatory system. Eukaryot. Cell 5, 696–711 (2006) | Article | PubMed | ChemPort |
14. Hutchings, N. R., Donelson, J. E. & Hill, K. L. Trypanin is a cytoskeletal linker protein and is required for cell motility in African trypanosomes. J. Cell Biol. 156, 867–877 (2002) | Article | PubMed | ChemPort |
15. Rupp, G. & Porter, M. E. A subunit of the dynein regulatory complex in Chlamydomonas is a homologue of a growth arrest-specific gene product. J. Cell Biol. 162, 47–57 (2003) | Article | PubMed | ChemPort |
16. Baron, D. M., Ralston, K. S., Kabututu, Z. P. & Hill, K. L. Functional genomics in Trypanosoma brucei identifies evolutionarily conserved components of motile flagella. J. Cell Sci. 120, 478–491 (2007) | Article | PubMed | ChemPort |
17. Hill, K. L., Hutchings, N. R., Grandgenett, P. M. & Donelson, J. E. T lymphocyte triggering factor of African trypanosomes is associated with the flagellar fraction of the cytoskeleton and represents a new family of proteins that are present in several divergent eukaryotes. J. Biol. Chem. 275, 39369–39378 (2000) | Article | PubMed | ChemPort |
18. Ralston, K. S. & Hill, K. L. Trypanin, a component of the flagellar dynein regulatory complex, is essential in bloodstream form African trypanosomes. PLoS Pathogens 2 doi:10.1371/journal.ppat.0020101 (2006) | Article | PubMed | ChemPort |
19. Bekker, J. M. et al. Direct interaction of Gas11 with microtubules: Implications for the dynein regulatory complex. Cell Motil. Cytoskeleton 64, 461–473 (2007) | Article | PubMed | ChemPort |
20. Colantonio, J. R. et al. Expanding the role of the dynein regulatory complex to non-axonemal functions: association of GAS11 with the golgi apparatus. Traffic 7, 538–548 (2006) | Article | PubMed | ChemPort |
21. Yeh, S. D. et al. Isolation and properties of Gas8, a growth arrest-specific gene regulated during male gametogenesis to produce a protein associated with the sperm motility apparatus. J. Biol. Chem. 277, 6311–6317 (2002) | Article | PubMed | ChemPort |
22. Whitmore, S. A. et al. Characterization and screening for mutations of the growth arrest-specific 11 (GAS11) and C16orf3 genes at 16q24.3 in breast cancer. Genomics 52, 325–331 (1998) | Article | PubMed | ChemPort |
23. Bok, J., Brunet, L. J., Howard, O., Burton, Q. & Wu, D. K. Role of hindbrain in inner ear morphogenesis: analysis of Noggin knockout mice. Dev. Biol. 311, 69–78 (2007) | Article | PubMed | ChemPort |
24. Leger, S. & Brand, M. Fgf8 and Fgf3 are required for zebrafish ear placode induction, maintenance and inner ear patterning. Mech. Dev. 119, 91–108 (2002) | Article | PubMed | ISI | ChemPort |
25. Mowbray, C., Hammerschmidt, M. & Whitfield, T. T. Expression of BMP signalling pathway members in the developing zebrafish inner ear and lateral line. Mech. Dev. 108, 179–184 (2001) | Article | PubMed | ChemPort |
26. Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullman, B. & Schilling, T. F. Stages of embryonic development of the zebrafish. Dev. Dyn. 203, 253–310 (1995) | PubMed | ISI | ChemPort |
27. Sullivan-Brown, J. et al. Zebrafish mutations affecting cilia motility share similar cystic phenotypes and suggest a mechanism of cyst formation that differs from pkd2 morphants. Dev. Biol. 314, 261–275 (2008) | Article | PubMed | ChemPort |
28. van Rooijen, E. et al. LRRC50, a conserved ciliary protein implicated in polycystic kidney disease. J. Am. Soc. Nephrol. 19, 1128–1138 (2008) | Article | PubMed | ChemPort |
29. Kawakami, Y., Raya, A., Raya, R. M., Rodriguez-Esteban, C. & Belmonte, J. C. Retinoic acid signalling links left–right asymmetric patterning and bilaterally symmetric somitogenesis in the zebrafish embryo. Nature 435, 165–171 (2005) | Article | PubMed | ISI | ChemPort |
30. Shastry, B. S. Mammalian cochlear genes and hereditary deafness. Microb. Comp. Genomics 5, 61–69 (2000) | Article | PubMed | ChemPort |
31. Westerfield, M. The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio rerio) (Univ. Oregon Press, 1993)
32. Curwen, V. et al. The Ensembl automatic gene annotation system. Genome Res. 14, 942–950 (2004) | Article | PubMed | ISI | ChemPort |
33. Chen, J. N. & Fishman, M. C. Zebrafish tinman homolog demarcates the heart field and initiates myocardial differentiation. Development 122, 3809–3816 (1996) | PubMed | ISI | ChemPort |
34. Drees, B. L. et al. A protein interaction map for cell polarity development. J. Cell Biol. 154, 549–576 (2001) | Article | PubMed | ISI | ChemPort |

Supplementary Information

Supplementary information accompanies this paper.

Acknowledgements

We are grateful to R. Crosbie for discussions and encouragement throughout the course of the project. We thank I. Drummond and C. Nguyen for discussions and comments on the work. We are grateful to L. Trinh, O. Bricaud and A. Collazo for sharing reagents and for providing probes, as well as to all the members of the Fraser laboratory for discussions, in particular M. Liebling and W. Supatto for sharing Matlab scripts and comments. We are grateful to Z. P. Kabututu for performing the lrrc50 reverse transcriptase PCR experiments. J.V. was supported by a fellowship from the Human Frontier Science Program, D.W. was supported by the NIH Medical Scientist Training Program at UCLA/Caltech. J.R.C. was supported by NIH RSDA training grant no. M07185 and a Warsaw Fellowship from the MIMG Department at UCLA. A.D.L is supported by an NSF fellowship. This work was supported by NIH grant R01 HL081799 (J.C.), NIH grant R01AI52348 and Beckman Young Investigator Award (K.L.H.).

Author Contributions J.R.C., J.V., D.W. and K.L.H. designed the experiments and interpreted the results. J.R.C, J.V. and D.W. conducted the experiments. J.V. and D.W. developed the equipment and systems for and performed in vivo video imaging for the quantitative flow study and analyzed the data with J.R.C., S.F. and K.L.H. A.D.L. assisted with in situ hybridization. A.D.L. and J.C. provided guidance on gas8 morpholino injections. The manuscript was written by J.R.C., J.V., D.W. and K.L.H. All authors discussed the results and commented on the manuscript.

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