Article | Published:

Myosin-Va is required for preciliary vesicle transportation to the mother centriole during ciliogenesis

Nature Cell Biologyvolume 20pages175185 (2018) | Download Citation


Primary cilia play essential roles in signal transduction and development. The docking of preciliary vesicles at the distal appendages of a mother centriole is an initial/critical step of ciliogenesis, but the mechanisms are unclear. Here, we demonstrate that myosin-Va mediates the transportation of preciliary vesicles to the mother centriole and reveal the underlying mechanism. We also show that the myosin-Va-mediated transportation of preciliary vesicles is the earliest event that defines the onset of ciliogenesis. Depletion of myosin-Va significantly inhibits the attachment of preciliary vesicles to the distal appendages of the mother centriole and decreases cilia assembly. Myosin-Va functions upstream of EHD1- and Rab11-mediated ciliary vesicle formation. Importantly, dynein mediates myosin-Va-associated preciliary vesicle transportation to the pericentrosomal region along microtubules, while myosin-Va mediates preciliary vesicle transportation from the pericentrosomal region to the distal appendages of the mother centriole via the Arp2/3-associated branched actin network.

Access optionsAccess options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Additional information

Publisher’s note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.


  1. 1.

    Gerdes, J. M., Davis, E. E. & Katsanis, N. The vertebrate primary cilium in development, homeostasis, and disease. Cell 137, 32–45 (2009).

  2. 2.

    Nigg, E. A. & Raff, J. W. Centrioles, centrosomes, and cilia in health and disease. Cell 139, 663–678 (2009).

  3. 3.

    Goetz, S. C. & Anderson, K. V. The primary cilium: a signalling centre during vertebrate development. Nat. Rev. Genet. 11, 331–344 (2010).

  4. 4.

    Hildebrandt, F., Benzing, T. & Katsanis, N. Ciliopathies. N. Engl. J. Med. 364, 1533–1543 (2011).

  5. 5.

    Sung, C. H. & Leroux, M. R. The roles of evolutionarily conserved functional modules in cilia-related trafficking. Nat. Cell Biol. 15, 1387–1397 (2013).

  6. 6.

    Sanchez, I. & Dynlacht, B. D. Cilium assembly and disassembly. Nat. Cell Biol. 18, 711–717 (2016).

  7. 7.

    Sorokin, S. P. Reconstructions of centriole formation and ciliogenesis in mammalian lungs. J. Cell Sci. 3, 207–230 (1968).

  8. 8.

    Lu, Q. et al. Early steps in primary cilium assembly require EHD1/EHD3-dependent ciliary vesicle formation. Nat. Cell Biol. 17, 228–240 (2015).

  9. 9.

    Yoshimura, S., Egerer, J., Fuchs, E., Haas, A. K. & Barr, F. A. Functional dissection of Rab GTPases involved in primary cilium formation. J. Cell Biol. 178, 363–369 (2007).

  10. 10.

    Nachury, M. V. et al. A core complex of BBS proteins cooperates with the GTPase Rab8 to promote ciliary membrane biogenesis. Cell 129, 1201–1213 (2007).

  11. 11.

    Knodler, A. et al. Coordination of Rab8 and Rab11 in primary ciliogenesis. Proc. Natl Acad. Sci. USA 107, 6346–6351 (2010).

  12. 12.

    Westlake, C. J. et al. Primary cilia membrane assembly is initiated by Rab11 and transport protein particle II (TRAPPII) complex-dependent trafficking of Rabin8 to the centrosome. Proc. Natl Acad. Sci. USA 108, 2759–2764 (2011).

  13. 13.

    Feng, S. et al. A Rab8 guanine nucleotide exchange factor–effector interaction network regulates primary ciliogenesis. J. Biol. Chem. 287, 15602–15609 (2012).

  14. 14.

    Chiba, S., Amagai, Y., Homma, Y., Fukuda, M. & Mizuno, K. NDR2-mediated Rabin8 phosphorylation is crucial for ciliogenesis by switching binding specificity from phosphatidylserine to Sec15. EMBO J. 32, 874–885 (2013).

  15. 15.

    Spektor, A., Tsang, W. Y., Khoo, D. & Dynlacht, B. D. Cep97 and CP110 suppress a cilia assembly program. Cell 130, 678–690 (2007).

  16. 16.

    Woolner, S. & Bement, W. M. Unconventional myosins acting unconventionally. Trends Cell Biol. 19, 245–252 (2009).

  17. 17.

    Hammer, J. A. III & Sellers, J. R. Walking to work: roles for class V myosins as cargo transporters. Nat. Rev. Mol. Cell Biol. 13, 13–26 (2012).

  18. 18.

    Jin, Y. et al. Myosin V transports secretory vesicles via a Rab GTPase cascade and interaction with the exocyst complex. Dev. Cell 21, 1156–1170 (2011).

  19. 19.

    Santiago-Tirado, F. H., Legesse-Miller, A., Schott, D. & Bretscher, A. PI4P and Rab inputs collaborate in myosin-V-dependent transport of secretory compartments in yeast. Dev. Cell 20, 47–59 (2011).

  20. 20.

    Donovan, K. W. & Bretscher, A. Myosin-V is activated by binding secretory cargo and released in coordination with Rab/exocyst function. Dev. Cell 23, 769–781 (2012).

  21. 21.

    Donovan, K. W. & Bretscher, A. Tracking individual secretory vesicles during exocytosis reveals an ordered and regulated process. J. Cell Biol. 210, 181–189 (2015).

  22. 22.

    Lindsay, A. J. et al. Identification and characterization of multiple novel Rab-myosin Va interactions. Mol. Biol. Cell 24, 3420–3434 (2013).

  23. 23.

    Ali, M. Y., Lu, H., Bookwalter, C. S., Warshaw, D. M. & Trybus, K. M. Myosin V and kinesin act as tethers to enhance each others’ processivity. Proc. Natl Acad. Sci. USA 105, 4691–4696 (2008).

  24. 24.

    Wada, F. et al. Myosin Va and endoplasmic reticulum calcium channel complex regulates membrane export during axon guidance. Cell Rep. 15, 1329–1344 (2016).

  25. 25.

    Assis, L. H. et al. The molecular motor myosin Va interacts with the cilia-centrosomal protein RPGRIP1L. Sci. Rep. 7, 43692 (2017).

  26. 26.

    Kohli, P. et al. The ciliary membrane-associated proteome reveals actin-binding proteins as key components of cilia. EMBO Rep. 18, 1521–1535 (2017).

  27. 27.

    Hong, H., Kim, J. & Kim, J. Myosin heavy chain 10 (MYH10) is required for centriole migration during the biogenesis of primary cilia. Biochem. Biophys. Res. Commun. 461, 180–185 (2015).

  28. 28.

    Franco, I. et al. Phosphoinositide 3-kinase-C2α regulates polycystin-2 ciliary entry and protects against kidney cyst formation. J. Am. Soc. Nephrol. 27, 1135–1144 (2016).

  29. 29.

    Franco, I. et al. PI3K class II alpha controls spatially restricted endosomal PtdIns3P and Rab11 activation to promote primary cilium function. Dev. Cell 28, 647–658 (2014).

  30. 30.

    Sorokin, S. Centrioles and the formation of rudimentary cilia by fibroblasts and smooth muscle cells. J. Cell Biol. 15, 363–377 (1962).

  31. 31.

    Ghossoub, R., Molla-Herman, A., Bastin, P. & Benmerah, A. The ciliary pocket: a once-forgotten membrane domain at the base of cilia. Biol. Cell 103, 131–144 (2011).

  32. 32.

    Molla-Herman, A. et al. The ciliary pocket: an endocytic membrane domain at the base of primary and motile cilia. J. Cell Sci. 123, 1785–1795 (2010).

  33. 33.

    Graser, S. et al. Cep164, a novel centriole appendage protein required for primary cilium formation. J. Cell Biol. 179, 321–330 (2007).

  34. 34.

    Schmidt, K. N. et al. Cep164 mediates vesicular docking to the mother centriole during early steps of ciliogenesis. J. Cell Biol. 199, 1083–1101 (2012).

  35. 35.

    Joo, K. et al. CCDC41 is required for ciliary vesicle docking to the mother centriole. Proc. Natl Acad. Sci. USA 110, 5987–5992 (2013).

  36. 36.

    Tanos, B. E. et al. Centriole distal appendages promote membrane docking, leading to cilia initiation. Genes Dev. 27, 163–168 (2013).

  37. 37.

    Tsakraklides, V. et al. Subcellular localization of GFP-myosin-V in live mouse melanocytes. J. Cell Sci. 112, 2853–2865 (1999).

  38. 38.

    Tai, A. W., Chuang, J. Z., Bode, C., Wolfrum, U. & Sung, C. H. Rhodopsin’s carboxy-terminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 97, 877–887 (1999).

  39. 39.

    Tai, A. W., Chuang, J. Z. & Sung, C. H. Cytoplasmic dynein regulation by subunit heterogeneity and its role in apical transport. J. Cell Biol. 153, 1499–1509 (2001).

  40. 40.

    Palmer, K. J., Hughes, H. & Stephens, D. J. Specificity of cytoplasmic dynein subunits in discrete membrane-trafficking steps. Mol. Biol. Cell 20, 2885–2899 (2009).

  41. 41.

    Firestone, A. J. et al. Small-molecule inhibitors of the AAA+ ATPase motor cytoplasmic dynein. Nature 484, 125–129 (2012).

  42. 42.

    Yadav, S., Puthenveedu, M. A. & Linstedt, A. D. Golgin160 recruits the dynein motor to position the Golgi apparatus. Dev. Cell 23, 153–165 (2012).

  43. 43.

    Quintyne, N. J. & Schroer, T. A. Distinct cell cycle-dependent roles for dynactin and dynein at centrosomes. J. Cell Biol. 159, 245–254 (2002).

  44. 44.

    Farina, F. et al. The centrosome is an actin-organizing centre. Nat. Cell Biol. 18, 65–75 (2016).

  45. 45.

    Kolega, J., Janson, L. W. & Taylor, D. L. The role of solation–contraction coupling in regulating stress fiber dynamics in nonmuscle cells. J. Cell Biol. 114, 993–1003 (1991).

  46. 46.

    Kim, J. et al. Functional genomic screen for modulators of ciliogenesis and cilium length. Nature 464, 1048–1051 (2010).

  47. 47.

    Kim, J. et al. Actin remodelling factors control ciliogenesis by regulating YAP/TAZ activity and vesicle trafficking. Nat. Commun. 6, 6781 (2015).

  48. 48.

    Chang, C. W., Hsu, W. B., Tsai, J. J., Tang, C. J. & Tang, T. K. CEP295 interacts with microtubules and is required for centriole elongation. J. Cell Sci. 129, 2501–2513 (2016).

  49. 49.

    Chen, H. Y. et al. Human microcephaly protein RTTN interacts with STIL and is required to build full-length centrioles. Nat. Commun. 8, 247 (2017).

  50. 50.

    Mali, P. et al. RNA-guided human genome engineering via Cas9. Science 339, 823–826 (2013).

  51. 51.

    Wang, W. J. et al. De novo centriole formation in human cells is error-prone and does not require SAS-6 self-assembly. eLife 4, e10586 (2015).

  52. 52.

    Demmerle, J. et al. Strategic and practical guidelines for successful structured illumination microscopy. Nat. Protoc. 12, 988–1010 (2017).

  53. 53.

    Ball, G. et al. SIMcheck: a toolbox for successful super-resolution structured illumination microscopy. Sci. Rep. 5, 15915 (2015).

  54. 54.

    Reddick, L. E. & Alto, N. M. Correlative light and electron microscopy (CLEM) as a tool to visualize microinjected molecules and their eukaryoticsub-cellular targets. J. Vis. Exp. 4, e3650 (2012).

Download references


The authors acknowledge support from the sequencing core facility (IBMS), the confocal imaging core facilities (IBMS, IMB, NPAS) and the EM core facilities (IMB, ICOB) of Academia Sinica. This work was supported by grants from the Ministry of Science and Technology, Taiwan (MOST 105-2321-B001-016) and the Academia Sinica Investigator Award.

Author information


  1. Taiwan International Graduate Program in Interdisciplinary Neuroscience, National Yang-Ming University and Academia Sinica, Taipei, Taiwan

    • Chien-Ting Wu
    •  & Tang K. Tang
  2. Institute of Biomedical Sciences, Academia Sinica, Taipei, Taiwan

    • Chien-Ting Wu
    • , Hsin-Yi Chen
    •  & Tang K. Tang


  1. Search for Chien-Ting Wu in:

  2. Search for Hsin-Yi Chen in:

  3. Search for Tang K. Tang in:


C.-T.W., a PhD student at the Taiwan International Graduate Program in Interdisciplinary Neuroscience, National Yang-Ming University and Academia Sinica, performed most of the experiments, designed the study, interpreted data and wrote the initial draft of the manuscript. H.-Y.C. performed experiments. T.K.T. conceived and designed the study, interpreted the data and wrote the manuscript.

Competing interests

The authors declare no competing financial interests.

Corresponding author

Correspondence to Tang K. Tang.

Supplementary information

  1. Supplementary Information

    Supplementary Figures 1–8, Supplementary legends.

  2. Life Sciences Reporting Summary

  3. Supplementary Table 1

    List of antibodies and dilutions used in this study.

  4. Supplementary Table 2

    List of siRNA sequences used in this study.


  1. Supplementary Video 1

    Related to Supplementary Fig. 2b. RPE1-based inducible cells expressing GFP-Myo-Va-GTD and mCherry-Arl13b were treated with DOX for 24 h and then serum-starved. Live cell images were taken using a LSM780 Carl Zeiss confocal system under controlled CO2 (5%) and temperature (37 °C).

  2. Supplementary Video 2

    Related to Supplementary Fig. 2d. NIH3T3-based inducible cells expressing GFP-Myo-Va-GTD and mCherry-Arl13b were treated with DOX for 24 h and then serum-starved. Live cell images were taken using a LSM780 Carl Zeiss confocal system under controlled CO2 (5%) and temperature (37 °C).

  3. Supplementary Video 3

    Related to Fig. 2f. IMCD3-based inducible cells expressing GFP-Myo-Va-GTD and mCherry-Arl13b were treated with DOX for 24 h and then serum-starved. Live cell images were taken using a LSM780 Carl Zeiss confocal system under controlled CO2 (5%) and temperature (37 °C).

  4. Supplementary Video 4

    Related to Supplementary Fig. 1e. RPE1-based inducible cells expressing GFP-EHD1 and mCherry-Myo-Va-GTD were treated with DOX for 24 h and then serum-starved. Live cell images were taken using a LSM780 Carl Zeiss confocal system under controlled CO2 (5%) and temperature (37 °C).

About this article

Publication history