Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

Regulation of microtubule dynamics, mechanics and function through the growing tip

Nature Reviews Molecular Cell Biology volume 22, pages 777–795 (2021)Cite this article

Subjects

Abstract

Microtubule dynamics and their control are essential for the normal function and division of all eukaryotic cells. This plethora of functions is, in large part, supported by dynamic microtubule tips, which can bind to various intracellular targets, generate mechanical forces and couple with actin microfilaments. Here, we review progress in the understanding of microtubule assembly and dynamics, focusing on new information about the structure of microtubule tips. First, we discuss evidence for the widely accepted GTP cap model of microtubule dynamics. Next, we address microtubule dynamic instability in the context of structural information about assembly intermediates at microtubule tips. Three currently discussed models of microtubule assembly and dynamics are reviewed. These are considered in the context of established facts and recent data, which suggest that some long-held views must be re-evaluated. Finally, we review structural observations about the tips of microtubules in cells and describe their implications for understanding the mechanisms of microtubule regulation by associated proteins, by mechanical forces and by microtubule-targeting drugs, prominently including cancer chemotherapeutics.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: The multiple roles of microtubule dynamics.
Fig. 2: Reorganizations of the microtubule cytoskeleton that rely on tubulin polymer dynamics.
Fig. 3: Current models for microtubule assembly and dynamics.
Fig. 4: Microtubule dynamics controlled by associated proteins, drugs and mechanical force.

Similar content being viewed by others

References

  1. Mitchison, T. & Kirschner, M. Dynamic instability of microtubule growth. Nature 312, 237–242 (1984). This paper presents data on microtubule dynamics in vitro that led to the formation of the theory of dynamic instability.

    CAS  PubMed  Google Scholar 

  2. Janke, C. & Magiera, M. M. The tubulin code and its role in controlling microtubule properties and functions. Nat. Rev. Mol. Cell Biol. 21, 307–326 (2020).

    CAS  PubMed  Google Scholar 

  3. Goodson, H. V. & Jonasson, E. M. Microtubules and microtubule-associated proteins. Cold Spring Harb. Perspect. Biol. 10, a022608 (2018).

    PubMed  PubMed Central  Google Scholar 

  4. Lomakin, A. J. et al. CLIP-170-dependent capture of membrane organelles by microtubules initiates minus-end directed transport. Dev. Cell 17, 323–333 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  5. Lomakin, A. J. et al. Stimulation of the CLIP-170-dependent capture of membrane organelles by microtubules through fine tuning of microtubule assembly dynamics. Mol. Biol. Cell 22, 4029–4037 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  6. Kanfer, G. et al. CENP-F couples cargo to growing and shortening microtubule ends. Mol. Biol. Cell 28, 2400–2409 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Kirschner, M. W. & Mitchison, T. Microtubule dynamics. Nature 324, 621 (1986).

    CAS  PubMed  Google Scholar 

  8. Holy, T. E. & Leibler, S. Dynamic instability of microtubules as an efficient way to search in space. Proc. Natl Acad. Sci. USA 91, 5682–5685 (1994).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. Zaytsev, A. V. & Grishchuk, E. L. Basic mechanism for biorientation of mitotic chromosomes is provided by the kinetochore geometry and indiscriminate turnover of kinetochore microtubules. Mol. Biol. Cell 26, 3985–3998 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  10. Wollman, R. et al. Efficient chromosome capture requires a bias in the ‘search-and-capture’ process during mitotic-spindle assembly. Curr. Biol. 15, 828–832 (2005).

    CAS  PubMed  Google Scholar 

  11. Dogterom, M. & Koenderink, G. H. Actin–microtubule crosstalk in cell biology. Nat. Rev. Mol. Cell Biol. 20, 38–54 (2019).

    CAS  PubMed  Google Scholar 

  12. Seetharaman, S. & Etienne-Manneville, S. Cytoskeletal crosstalk in cell migration. Trends Cell Biol. 30, 720–735 (2020).

    CAS  PubMed  Google Scholar 

  13. Jiang, K. et al. A proteome-wide screen for mammalian SxIP motif-containing microtubule plus-end tracking proteins. Curr. Biol. 22, 1800–1807 (2012).

    CAS  PubMed  Google Scholar 

  14. Kodama, A., Karakesisoglou, I., Wong, E., Vaezi, A. & Fuchs, E. ACF7: an essential integrator of microtubule dynamics. Cell 115, 343–354 (2003).

    CAS  PubMed  Google Scholar 

  15. Stroud, M. J. et al. GAS2-like proteins mediate communication between microtubules and actin through interactions with end-binding proteins. J. Cell Sci. 127, 2672–2682 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  16. Henty-Ridilla, J. L., Rankova, A., Eskin, J. A., Kenny, K. & Goode, B. L. Accelerated actin filament polymerization from microtubule plus ends. Science 352, 1004–1009 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  17. Daga, R. R., Yonetani, A. & Chang, F. Asymmetric microtubule pushing forces in nuclear centering. Curr. Biol. 16, 1544–1550 (2006).

    CAS  PubMed  Google Scholar 

  18. Tran, P. T., Marsh, L., Doye, V., Inoué, S. & Chang, F. A mechanism for nuclear positioning in fission yeast based on microtubule pushing. J. Cell Biol. 153, 397–412 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Zhao, T., Graham, O. S., Raposo, A. & Johnston, D. S. Growing microtubules push the oocyte nucleus to polarize the Drosophila dorsal–ventral axis. Science 336, 999–1003 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Yeh, E., Skibbens, R. V., Cheng, J. W., Salmon, E. D. & Bloom, K. Spindle dynamics and cell cycle regulation of dynein in the budding yeast, Saccharomyces cerevisiae. J. Cell Biol. 130, 687–700 (1995).

    CAS  PubMed  Google Scholar 

  21. Penfield, L. et al. Dynein pulling forces counteract lamin-mediated nuclear stability during nuclear envelope repair. MBoC 29, 852–868 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Gönczy, P., Pichler, S., Kirkham, M. & Hyman, A. A. Cytoplasmic dynein is required for distinct aspects of Mtoc positioning, including centrosome separation, in the one cell stage Caenorhabditis elegans embryo. J. Cell Biol. 147, 135–150 (1999).

    PubMed  PubMed Central  Google Scholar 

  23. Tanimoto, H., Kimura, A. & Minc, N. Shape–motion relationships of centering microtubule asters. J. Cell Biol. 212, 777–787 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Laan, L. et al. Cortical dynein controls microtubule dynamics to generate pulling forces that position microtubule asters. Cell 148, 502–514 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Tolic´-Nørrelykke, I. M., Sacconi, L., Thon, G. & Pavone, F. S. Positioning and elongation of the fission yeast spindle by microtubule-based pushing. Curr. Biol. 14, 1181–1186 (2004).

    PubMed  Google Scholar 

  26. Burakov, A., Nadezhdina, E., Slepchenko, B. & Rodionov, V. Centrosome positioning in interphase cells. J. Cell Biol. 162, 963–969 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Meaders, J. L., de Matos, S. N. & Burgess, D. R. A pushing mechanism for microtubule aster positioning in a large cell type. Cell Rep. 33, 108213 (2020).

    CAS  PubMed  Google Scholar 

  28. Tilney, L. G. & Porter, K. R. Studies on the microtubules in heliozoa. II. The effect of low temperature on these structures in the formation and maintenance of the axopodia. J. Cell Biol. 34, 327–343 (1967).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Tilney, L. G., Hiramoto, Y. & Marsland, D. Studies on the microtubules in heliozoa. 3. A pressure analysis of the role of these structures in the formation and maintenance of the axopodia of Actinosphaerium nucleofilum (Barrett). J. Cell Biol. 29, 77–95 (1966).

    CAS  PubMed  PubMed Central  Google Scholar 

  30. Brangwynne, C. P. et al. Microtubules can bear enhanced compressive loads in living cells because of lateral reinforcement. J. Cell Biol. 173, 733–741 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Rodríguez-García, R. et al. Mechanisms of motor-independent membrane remodeling driven by dynamic microtubules. Curr. Biol. 30, 972–987.e12 (2020). This paper clearly demonstrates the ability of microtubules to exert pulling and pushing forces on membranes in vitro.

    PubMed  PubMed Central  Google Scholar 

  32. Grishchuk, E. L. & McIntosh, J. R. Microtubule depolymerization can drive poleward chromosome motion in fission yeast. EMBO J. 25, 4888–4896 (2006). This paper provides the first direct evidence that microtubule depolymerization can generate force in living cells.

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Tanaka, K., Kitamura, E., Kitamura, Y. & Tanaka, T. U. Molecular mechanisms of microtubule-dependent kinetochore transport toward spindle poles. J. Cell Biol. 178, 269–281 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Vukušic´, K., Buda, R. & Tolic´, I. M. Force-generating mechanisms of anaphase in human cells. J. Cell Sci. 132, jcs231985 (2019).

    PubMed  Google Scholar 

  35. McIntosh, J. R. Anaphase A. Semin. Cell Dev. Biol. https://doi.org/10.1016/j.semcdb.2021.03.009 (2021).

    Article  PubMed  Google Scholar 

  36. Meiring, J. C. M., Shneyer, B. I. & Akhmanova, A. Generation and regulation of microtubule network asymmetry to drive cell polarity. Curr. Opin. Cell Biol. 62, 86–95 (2020).

    CAS  PubMed  Google Scholar 

  37. Heald, R. & Khodjakov, A. Thirty years of search and capture: the complex simplicity of mitotic spindle assembly. J. Cell Biol. 211, 1103–1111 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. Röper, K. Microtubules enter centre stage for morphogenesis. Philos. Trans. R. Soc. B https://doi.org/10.1098/rstb.2019.0557 (2020).

    Article  Google Scholar 

  39. Kaverina, I., Rottner, K. & Small, J. V. Targeting, capture, and stabilization of microtubules at early focal adhesions. J. Cell Biol. 142, 181–190 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Stehbens, S. J. et al. CLASPs link focal adhesion-associated microtubule capture to localized exocytosis and adhesion site turnover. Nat. Cell Biol. 16, 561–573 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Kopf, A. et al. Microtubules control cellular shape and coherence in amoeboid migrating cells. J. Cell Biol. 219, e201907154 (2020).

    PubMed  PubMed Central  Google Scholar 

  42. Tabdanov, E. D. et al. Engineering T cells to enhance 3D migration through structurally and mechanically complex tumor microenvironments. Nat. Commun. 12, 2815 (2021).

    CAS  PubMed  PubMed Central  Google Scholar 

  43. Schelski, M. & Bradke, F. Neuronal polarization: from spatiotemporal signaling to cytoskeletal dynamics. Mol. Cell. Neurosci. 84, 11–28 (2017).

    CAS  PubMed  Google Scholar 

  44. Witte, H., Neukirchen, D. & Bradke, F. Microtubule stabilization specifies initial neuronal polarization. J. Cell Biol. 180, 619–632 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Dupraz, S. et al. RhoA controls axon extension independent of specification in the developing brain. Curr. Biol. 29, 3874–3886.e9 (2019).

    CAS  PubMed  Google Scholar 

  46. Singh, A. et al. Polarized microtubule dynamics directs cell mechanics and coordinates forces during epithelial morphogenesis. Nat. Cell Biol. 20, 1126–1133 (2018).

    CAS  PubMed  Google Scholar 

  47. Patel-Hett, S. et al. Visualization of microtubule growth in living platelets reveals a dynamic marginal band with multiple microtubules. Blood 111, 4605–4616 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. Diagouraga, B. et al. Motor-driven marginal band coiling promotes cell shape change during platelet activation. J. Cell Biol. 204, 177–185 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. Yi, J. et al. Centrosome repositioning in T cells is biphasic and driven by microtubule end-on capture-shrinkage. J. Cell Biol. 202, 779–792 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  50. Hooikaas, P. J. et al. Kinesin-4 KIF21B limits microtubule growth to allow rapid centrosome polarization in T cells. eLife 9, e62876 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  51. McIntosh, J. R., Grishchuk, E. L. & West, R. R. Chromosome–microtubule interactions during mitosis. Annu. Rev. Cell Dev. Biol. 18, 193–219 (2002).

    CAS  PubMed  Google Scholar 

  52. Kapoor, T. M. Metaphase spindle assembly. Biology 6, 8 (2017).

    PubMed Central  Google Scholar 

  53. Hyman, A. A., Salser, S., Drechsel, D. N., Unwin, N. & Mitchison, T. J. Role of GTP hydrolysis in microtubule dynamics: information from a slowly hydrolyzable analogue, GMPCPP. Mol. Biol. Cell 3, 1155–1167 (1992).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Walker, R. A., Inoué, S. & Salmon, E. D. Asymmetric behavior of severed microtubule ends after ultraviolet-microbeam irradiation of individual microtubules in vitro. J. Cell Biol. 108, 931–937 (1989).

    CAS  PubMed  Google Scholar 

  55. Tran, P. T., Walker, R. A. & Salmon, E. D. A metastable intermediate state of microtubule dynamic instability that differs significantly between plus and minus ends. J. Cell Biol. 138, 105–117 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  56. Caplow, M. & Shanks, J. Evidence that a single monolayer tubulin-GTP cap is both necessary and sufficient to stabilize microtubules. MBoC 7, 663–675 (1996).

    CAS  PubMed  PubMed Central  Google Scholar 

  57. Walker, R. A., Pryer, N. K. & Salmon, E. D. Dilution of individual microtubules observed in real time in vitro: evidence that cap size is small and independent of elongation rate. J. Cell Biol. 114, 73–81 (1991).

    CAS  PubMed  Google Scholar 

  58. Duellberg, C., Cade, N. I., Holmes, D. & Surrey, T. The size of the EB cap determines instantaneous microtubule stability. eLife 5, e13470 (2016). This paper presents important measurements of the size of the microtubule-stabilizing cap in vitro.

    PubMed  PubMed Central  Google Scholar 

  59. Maurer, S. P., Bieling, P., Cope, J., Hoenger, A. & Surrey, T. GTPγS microtubules mimic the growing microtubule end structure recognized by end-binding proteins (EBs). Proc. Natl Acad. Sci. USA 108, 3988–3993 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. Dimitrov, A. et al. Detection of GTP-tubulin conformation in vivo reveals a role for GTP remnants in microtubule rescues. Science 322, 1353–1356 (2008).

    CAS  PubMed  Google Scholar 

  61. Roostalu, J. et al. The speed of GTP hydrolysis determines GTP cap size and controls microtubule stability. eLife 9, e51992 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. Mandelkow, E. M., Mandelkow, E. & Milligan, R. A. Microtubule dynamics and microtubule caps: a time-resolved cryo-electron microscopy study. J. Cell Biol. 114, 977–991 (1991). This paper presents the first cryo-EM study of dynamic microtubules. The images help to define the allosteric model for nucleotide regulation of tubulin’s shape and, therefore, its polymerization dynamics.

    CAS  PubMed  Google Scholar 

  63. Marantz, R. & Shelanski, M. L. Structure of microtubular crystals induced by vinblastine in vitro. J. Cell Biol. 44, 234–238 (1970).

    CAS  PubMed  PubMed Central  Google Scholar 

  64. Müller-Reichert, T., Chrétien, D., Severin, F. & Hyman, A. A. Structural changes at microtubule ends accompanying GTP hydrolysis: information from a slowly hydrolyzable analogue of GTP, guanylyl (α,β)methylenediphosphonate. Proc. Natl Acad. Sci. USA 95, 3661–3666 (1998).

    PubMed  PubMed Central  Google Scholar 

  65. Wang, H.-W. & Nogales, E. Nucleotide-dependent bending flexibility of tubulin regulates microtubule assembly. Nature 435, 911–915 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. Howes, S. C. et al. Structural and functional differences between porcine brain and budding yeast microtubules. Cell Cycle 17, 278–287 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Ayukawa, R. et al. GTP-dependent formation of straight tubulin oligomers leads to microtubule nucleation. J. Cell Biol. 220, e202007033 (2021).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. McIntosh, J. R. et al. Microtubules grow by the addition of bent guanosine triphosphate tubulin to the tips of curved protofilaments. J. Cell Biol. 217, 2691–2708 (2018). This paper presents data on the shapes of microtubule tips growing either in cells or in vitro, as seen by electron tomography, using fast-frozen/freeze-substitution fixed cells and both fast-frozen and fixed samples for cryo-ET of frozen hydrated samples in vitro.

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Buey, R. M., Díaz, J. F. & Andreu, J. M. The nucleotide switch of tubulin and microtubule assembly: a polymerization-driven structural change. Biochemistry 45, 5933–5938 (2006).

    CAS  PubMed  Google Scholar 

  70. Aldaz, H., Rice, L. M., Stearns, T. & Agard, D. A. Insights into microtubule nucleation from the crystal structure of human γ-tubulin. Nature 435, 523–527 (2005).

    CAS  PubMed  Google Scholar 

  71. Nawrotek, A., Knossow, M. & Gigant, B. The determinants that govern microtubule assembly from the atomic structure of GTP-tubulin. J. Mol. Biol. 412, 35–42 (2011).

    CAS  PubMed  Google Scholar 

  72. Pecqueur, L. et al. A designed ankyrin repeat protein selected to bind to tubulin caps the microtubule plus end. Proc. Natl Acad. Sci. USA 109, 12011–12016 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. Brouhard, G. J. & Rice, L. M. The contribution of αβ-tubulin curvature to microtubule dynamics. J. Cell Biol. 207, 323–334 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  74. Gebremichael, Y., Chu, J.-W. & Voth, G. A. Intrinsic bending and structural rearrangement of tubulin dimer: molecular dynamics simulations and coarse-grained analysis. Biophys. J. 95, 2487–2499 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Grafmüller, A. & Voth, G. A. Intrinsic bending of microtubule protofilaments. Structure 19, 409–417 (2011).

    PubMed  Google Scholar 

  76. Igaev, M. & Grubmüller, H. Microtubule assembly governed by tubulin allosteric gain in flexibility and lattice induced fit. eLife 7, e34353 (2018).

    PubMed  PubMed Central  Google Scholar 

  77. Fedorov, V. A. et al. Mechanical properties of tubulin intra- and inter-dimer interfaces and their implications for microtubule dynamic instability. PLoS Comput. Biol. 15, e1007327 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. Tong, D. & Voth, G. A. Microtubule simulations provide insight into the molecular mechanism underlying dynamic instability. Biophys. J. 118, 2938–2951 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. Alushin, G. M. et al. High-resolution microtubule structures reveal the structural transitions in αβ-tubulin upon GTP hydrolysis. Cell 157, 1117–1129 (2014). This paper presents the first moderately high-resolution structure of the nucleotide-dependent microtubule lattices obtained with cryo-EM.

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Zhang, R., Alushin, G. M., Brown, A. & Nogales, E. Mechanistic origin of microtubule dynamic instability and its modulation by EB proteins. Cell 162, 849–859 (2015). This paper discusses the structure of tubulin in microtubules and how it is affected by external factors as seen by cryo-EM and image averaging.

    CAS  PubMed  PubMed Central  Google Scholar 

  81. Zhang, R., LaFrance, B. & Nogales, E. Separating the effects of nucleotide and EB binding on microtubule structure. Proc. Natl Acad. Sci. USA 115, E6191–E6200 (2018).

    PubMed  PubMed Central  Google Scholar 

  82. Manka, S. W. & Moores, C. A. The role of tubulin–tubulin lattice contacts in the mechanism of microtubule dynamic instability. Nat. Struct. Mol. Biol. 25, 607–615 (2018). This paper is an important structural study of tubulin–tubulin contacts in microtubule lattices, assembled in the presence of different nucleotides.

    CAS  PubMed  PubMed Central  Google Scholar 

  83. von Loeffelholz, O. et al. Nucleotide- and Mal3-dependent changes in fission yeast microtubules suggest a structural plasticity view of dynamics. Nat. Commun. 8, 2110 (2017).

    Google Scholar 

  84. Howes, S. C. et al. Structural differences between yeast and mammalian microtubules revealed by cryo-EM. J. Cell Biol. 216, 2669–2677 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Estévez-Gallego, J. et al. Structural model for differential cap maturation at growing microtubule ends. eLife 9, e50155 (2020).

    PubMed  PubMed Central  Google Scholar 

  86. Odde, D. J., Cassimeris, L. & Buettner, H. M. Kinetics of microtubule catastrophe assessed by probabilistic analysis. Biophys. J. 69, 796–802 (1995). This paper describes the discovery of the ‘microtubule ageing’ phenomenon.

    CAS  PubMed  PubMed Central  Google Scholar 

  87. Gardner, M. K., Zanic, M., Gell, C., Bormuth, V. & Howard, J. Depolymerizing kinesins Kip3 and MCAK shape cellular microtubule architecture by differential control of catastrophe. Cell 147, 1092–1103 (2011).

    CAS  PubMed  Google Scholar 

  88. Bowne-Anderson, H., Zanic, M., Kauer, M. & Howard, J. Microtubule dynamic instability: a new model with coupled GTP hydrolysis and multistep catastrophe. Bioessays 35, 452–461 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  89. Zakharov, P. et al. Molecular and mechanical causes of microtubule catastrophe and aging. Biophys. J. 109, 2574–2591 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. Coombes, C. E., Yamamoto, A., Kenzie, M. R., Odde, D. J. & Gardner, M. K. Evolving tip structures can explain age-dependent microtubule catastrophe. Curr. Biol. 23, 1342–1348 (2013). This paper uses fluorescence microscopy to study the elongation and ageing of microtubules in the presence of labelled tubulin.

    CAS  PubMed  PubMed Central  Google Scholar 

  91. Kirschner, M. W., Williams, R. C., Weingarten, M. & Gerhart, J. C. Microtubules from mammalian brain: some properties of their depolymerization products and a proposed mechanism of assembly and disassembly. Proc. Natl Acad. Sci. USA 71, 1159–1163 (1974).

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Simon, J. R. & Salmon, E. D. The structure of microtubule ends during the elongation and shortening phases of dynamic instability examined by negative-stain electron microscopy. J. Cell. Sci. 96, 571–582 (1990).

    CAS  PubMed  Google Scholar 

  93. VanBuren, V., Odde, D. J. & Cassimeris, L. Estimates of lateral and longitudinal bond energies within the microtubule lattice. Proc. Natl Acad. Sci. USA 99, 6035–6040 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  94. Chretien, D., Fuller, S. D. & Karsenti, E. Structure of growing microtubule ends: two-dimensional sheets close into tubes at variable rates. J. Cell Biol. 129, 1311–1328 (1995). This paper uses cryo-ET to provide an important description of the shapes of ends on microtubules elongating in vitro.

    CAS  PubMed  Google Scholar 

  95. Guesdon, A. et al. EB1 interacts with outwardly curved and straight regions of the microtubule lattice. Nat. Cell Biol. 18, 1102–1108 (2016). This paper is an extension of the work described by Chretien et al. (1995) that uses cryo-ET and immunolabelling to describe microtubule elongation.

    CAS  PubMed  Google Scholar 

  96. Stewman, S. F., Tsui, K. K. & Ma, A. Dynamic instability from non-equilibrium structural transitions on the energy landscape of microtubule. Cell Syst. 11, 608–624 (2020).

    CAS  PubMed  Google Scholar 

  97. Gudimchuk, N. B. et al. Mechanisms of microtubule dynamics and force generation examined with computational modeling and electron cryotomography. Nat. Commun. 11, 3765 (2020). This paper presents development and experimental testing of the model for microtubule growth through the straightening of curved protofilaments, as presented by McIntosh et al. (2018).

    PubMed  PubMed Central  Google Scholar 

  98. Atherton, J., Stouffer, M., Francis, F. & Moores, C. A. Microtubule architecture in vitro and in cells revealed by cryo-electron tomography. Acta Crystallogr. D 74, 572–584 (2018).

    CAS  Google Scholar 

  99. Margolin, G. et al. The mechanisms of microtubule catastrophe and rescue: implications from analysis of a dimer-scale computational model. Mol. Biol. Cell 23, 642–656 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  100. VandenBeldt, K. J. et al. Kinetochores use a novel mechanism for coordinating the dynamics of individual microtubules. Curr. Biol. 16, 1217–1223 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. McIntosh, J. R. et al. Conserved and divergent features of kinetochores and spindle microtubule ends from fibe species. J. Cell Biol. 200, 459–474 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. Zovko, S., Abrahams, J. P., Koster, A. J., Galjart, N. & Mommaas, A. M. Microtubule plus-end conformations and dynamics in the periphery of interphase mouse fibroblasts. Mol. Biol. Cell 19, 3138–3146 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. Hoog, J. L. et al. Electron tomography reveals a flared morphology on growing microtubule ends. J. Cell Sci. 124, 693–698 (2011).

    PubMed  PubMed Central  Google Scholar 

  104. Kukulski, W. et al. Correlated fluorescence and 3D electron microscopy with high sensitivity and spatial precision. J. Cell Biol. 192, 111–119 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Nehlig, A., Molina, A., Rodrigues-Ferreira, S., Honoré, S. & Nahmias, C. Regulation of end-binding protein EB1 in the control of microtubule dynamics. Cell Mol. Life Sci. 74, 2381–2393 (2017).

    CAS  PubMed  Google Scholar 

  106. Vitre, B. et al. EB1 regulates microtubule dynamics and tubulin sheet closure in vitro. Nat. Cell Biol. 10, 415–421 (2008).

    CAS  PubMed  Google Scholar 

  107. Komarova, Y. et al. Mammalian end binding proteins control persistent microtubule growth. J. Cell Biol. 184, 691–706 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Bieling, P. et al. Reconstitution of a microtubule plus-end tracking system in vitro. Nature 450, 1100–1105 (2007).

    CAS  PubMed  Google Scholar 

  109. Maurer, S. P. et al. EB1 accelerates two conformational transitions important for microtubule maturation and dynamics. Curr. Biol. 24, 372–384 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  110. Tirnauer, J. S., Grego, S., Salmon, E. D. & Mitchison, T. J. EB1–microtubule interactions in Xenopus egg extracts: role of EB1 in microtubule stabilization and mechanisms of targeting to microtubules. Mol. Biol. Cell 13, 3614–3626 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Maurer, S. P., Fourniol, F. J., Bohner, G., Moores, C. A. & Surrey, T. EBs recognize a nucleotide-dependent structural cap at growing microtubule ends. Cell 149, 371–382 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  112. Gardner, M. K. et al. Rapid microtubule self-assembly kinetics. Cell 146, 582–592 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. Gard, D. L. & Kirschner, M. W. A microtubule-associated protein from Xenopus eggs that specifically promotes assembly at the plus-end. J. Cell Biol. 105, 2203–2215 (1987).

    CAS  PubMed  Google Scholar 

  114. Akhmanova, A. et al. CLASPs are CLIP-115 and -170 associating proteins involved in the regional regulation of microtubule dynamics in motile fibroblasts. Cell 104, 923–935 (2001).

    CAS  PubMed  Google Scholar 

  115. Das, A., Dickinson, D. J., Wood, C. C., Goldstein, B. & Slep, K. C. Crescerin uses a TOG domain array to regulate microtubules in the primary cilium. Mol. Biol. Cell 26, 4248–4264 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  116. Tournebize, R. et al. Control of microtubule dynamics by the antagonistic activities of XMAP215 and XKCM1 in Xenopus egg extracts. Nat. Cell Biol. 2, 13–19 (2000).

    CAS  PubMed  Google Scholar 

  117. Gergely, F., Draviam, V. M. & Raff, J. W. The ch-TOG/XMAP215 protein is essential for spindle pole organization in human somatic cells. Genes Dev. 17, 336–341 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Garcia, M. A., Vardy, L., Koonrugsa, N. & Toda, T. Fission yeast ch-TOG/XMAP215 homologue Alp14 connects mitotic spindles with the kinetochore and is a component of the Mad2-dependent spindle checkpoint. EMBO J. 20, 3389–3401 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. Miller, M. P., Asbury, C. L. & Biggins, S. A TOG protein confers tension sensitivity to kinetochore–microtubule attachments. Cell 165, 1428 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  120. Al-Bassam, J. & Chang, F. Regulation of microtubule dynamics by TOG-domain proteins XMAP215/Dis1 and CLASP. Trends Cell Biol. 21, 604–614 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Slep, K. C. & Vale, R. D. Structural basis of microtubule plus end tracking by XMAP215, CLIP-170 and EB1. Mol. Cell 27, 976–991 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  122. Rice, L. M., Montabana, E. A. & Agard, D. A. The lattice as allosteric effector: structural studies of αβ- and γ-tubulin clarify the role of GTP in microtubule assembly. Proc. Natl Acad. Sci. USA 105, 5378–5383 (2008). This paper is the first clear statement of the ‘lattice model’ for how GTP-tubulin alters its properties to allow dynamic instability.

    CAS  PubMed  PubMed Central  Google Scholar 

  123. Fox, J. C., Howard, A. E., Currie, J. D., Rogers, S. L. & Slep, K. C. The XMAP215 family drives microtubule polymerization using a structurally diverse TOG array. MBoC 25, 2375–2392 (2014).

    PubMed  PubMed Central  Google Scholar 

  124. Byrnes, A. E. & Slep, K. C. TOG–tubulin binding specificity promotes microtubule dynamics and mitotic spindle formation. J. Cell Biol. 216, 1641–1657 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  125. Kinoshita, K., Arnal, I., Desai, A., Drechsel, D. N. & Hyman, A. A. Reconstitution of physiological microtubule dynamics using purified components. Science 294, 1340–1343 (2001).

    CAS  PubMed  Google Scholar 

  126. Brouhard, G. J. et al. XMAP215 is a processive microtubule polymerase. Cell 132, 79–88 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. Zanic, M., Widlund, P. O., Hyman, A. A. & Howard, J. Synergy between XMAP215 and EB1 increases microtubule growth rates to physiological levels. Nat. Cell Biol. 15, 688–693 (2013).

    CAS  PubMed  Google Scholar 

  128. Farmer, V., Arpag˘, G., Hall, S. & Zanic, M. XMAP215 promotes microtubule catastrophe by disrupting the growing microtubule end. Preprint at https://doi.org/10.1101/2020.12.29.424748 (2020).

  129. Ayaz, P. et al. A tethered delivery mechanism explains the catalytic action of a microtubule polymerase. eLife 3, e03069 (2014).

    PubMed  PubMed Central  Google Scholar 

  130. Drabek, K. et al. Role of CLASP2 in microtubule stabilization and the regulation of persistent motility. Curr. Biol. 16, 2259–2264 (2006).

    CAS  PubMed  Google Scholar 

  131. Sousa, A., Reis, R., Sampaio, P. & Sunkel, C. E. The Drosophila CLASP homologue, Mast/Orbit regulates the dynamic behaviour of interphase microtubules by promoting the pause state. Cell Motil. 64, 605–620 (2007).

    CAS  Google Scholar 

  132. Majumdar, S. et al. An isolated CLASP TOG domain suppresses microtubule catastrophe and promotes rescue. MBoC 29, 1359–1375 (2018).

    PubMed  PubMed Central  Google Scholar 

  133. Lawrence, E. J., Arpag˘, G., Norris, S. R. & Zanic, M. Human CLASP2 specifically regulates microtubule catastrophe and rescue. MBoC 29, 1168–1177 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  134. Mimori-Kiyosue, Y. et al. CLASP1 and CLASP2 bind to EB1 and regulate microtubule plus-end dynamics at the cell cortex. J. Cell Biol. 168, 141–153 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  135. Leano, J. B. & Slep, K. C. Structures of TOG1 and TOG2 from the human microtubule dynamics regulator CLASP1. PLoS ONE 14, e0219823 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  136. Aher, A. et al. CLASP suppresses microtubule catastrophes through a single TOG domain. Dev. Cell 46, 40–58.e8 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. Aher, A. et al. CLASP mediates microtubule repair by restricting lattice damage and regulating tubulin incorporation. Curr. Biol. 30, 2175–2183.e6 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  138. Howell, B., Larsson, N., Gullberg, M. & Cassimeris, L. Dissociation of the tubulin-sequestering and microtubule catastrophe-promoting activities of oncoprotein 18/stathmin. Mol. Biol. Cell 10, 105–118 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  139. Wang, C., Cormier, A., Gigant, B. & Knossow, M. Insight into the GTPase activity of tubulin from complexes with stathmin-like domains. Biochemistry 46, 10595–10602 (2007).

    CAS  PubMed  Google Scholar 

  140. Manning, A. L. et al. The kinesin-13 proteins Kif2a, Kif2b, and Kif2c/MCAK have distinct roles during mitosis in human cells. Mol. Biol. Cell 18, 2970–2979 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  141. Ganem, N. J., Godinho, S. A. & Pellman, D. A mechanism linking extra centrosomes to chromosomal instability. Nature 460, 278 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. Ohi, R., Burbank, K., Liu, Q. & Mitchison, T. J. Nonredundant functions of kinesin-13s during meiotic spindle assembly. Curr. Biol. 17, 953–959 (2007).

    CAS  PubMed  Google Scholar 

  143. Homma, N. et al. Kinesin superfamily protein 2A (KIF2A) functions in suppression of collateral branch extension. Cell 114, 229–239 (2003).

    CAS  PubMed  Google Scholar 

  144. Kobayashi, T., Tsang, W. Y., Li, J., Lane, W. & Dynlacht, B. D. Centriolar kinesin Kif24 interacts with CP110 to remodel microtubules and regulate ciliogenesis. Cell 145, 914–925 (2011).

    CAS  PubMed  Google Scholar 

  145. Wang, W. et al. Insight into microtubule disassembly by kinesin-13s from the structure of Kif2C bound to tubulin. Nat. Commun. 8, 70 (2017).

    PubMed  PubMed Central  Google Scholar 

  146. Trofimova, D. et al. Ternary complex of Kif2A-bound tandem tubulin heterodimers represents a kinesin-13-mediated microtubule depolymerization reaction intermediate. Nat. Commun. 9, 2628 (2018).

    PubMed  PubMed Central  Google Scholar 

  147. Paydar, M. & Kwok, B. H. Evidence for conformational change-induced hydrolysis of β-tubulin-GTP. Sneak Peak https://papers.ssrn.com/abstract=3687033 (2020)

  148. Helenius, J., Brouhard, G., Kalaidzidis, Y., Diez, S. & Howard, J. The depolymerizing kinesin MCAK uses lattice diffusion to rapidly target microtubule ends. Nature 441, 115–119 (2006).

    CAS  PubMed  Google Scholar 

  149. Oguchi, Y., Uchimura, S., Ohki, T., Mikhailenko, S. V. & Ishiwata, S. The bidirectional depolymerizer MCAK generates force by disassembling both microtubule ends. Nat. Cell Biol. 13, 846–852 (2011).

    CAS  PubMed  Google Scholar 

  150. Varga, V. et al. Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner. Nat. Cell Biol. 8, 957–962 (2006).

    CAS  PubMed  Google Scholar 

  151. Arellano-Santoyo, H. et al. A tubulin binding switch underlies Kip3/Kinesin-8 depolymerase activity. Dev. Cell 42, 37–51.e8 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Varga, V., Leduc, C., Bormuth, V., Diez, S. & Howard, J. Kinesin-8 motors act cooperatively to mediate length-dependent microtubule depolymerization. Cell 138, 1174–1183 (2009).

    CAS  PubMed  Google Scholar 

  153. Chen, G.-Y. et al. Kinesin-5 promotes microtubule nucleation and assembly by stabilizing a lattice-competent conformation of tubulin. Curr. Biol. 29, 2259–2269.e4 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  154. Ayaz, P., Ye, X., Huddleston, P., Brautigam, C. A. & Rice, L. M. A TOG:αβ-tubulin complex structure reveals conformation-based mechanisms for a microtubule polymerase. Science 337, 857–860 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Ravelli, R. B. G. et al. Insight into tubulin regulation from a complex with colchicine and a stathmin-like domain. Nature 428, 198–202 (2004).

    CAS  PubMed  Google Scholar 

  156. Gupta, K. K. et al. Mechanism for the catastrophe-promoting activity of the microtubule destabilizer Op18/stathmin. PNAS 110, 20449–20454 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  157. Jordan, M. A. & Wilson, L. Microtubules as a target for anticancer drugs. Nat. Rev. Cancer 4, 253–265 (2004).

    CAS  PubMed  Google Scholar 

  158. Brunden, K. R., Trojanowski, J. Q., Smith, A. B., Lee, V. M.-Y. & Ballatore, C. Microtubule-stabilizing agents as potential therapeutics for neurodegenerative disease. Bioorg. Med. Chem. 22, 5040–5049 (2014).

    CAS  PubMed  Google Scholar 

  159. Steinmetz, M. O. & Prota, A. E. Microtubule-targeting agents: strategies to hijack the cytoskeleton. Trends Cell Biol. 28, 776–792 (2018).

    CAS  PubMed  Google Scholar 

  160. Guo, H., Li, X., Guo, Y. & Zhen, L. An overview of tubulin modulators deposited in protein data bank. Med. Chem. Res. 28, 927–937 (2019).

    CAS  Google Scholar 

  161. Gallego-Jara, J., Lozano-Terol, G., Sola-Martínez, R. A., Cánovas-Díaz, M. & de Diego Puente, T. A. Compressive review about Taxol®: history and future challenges. Molecules 25, 5986 (2020).

    CAS  PubMed Central  Google Scholar 

  162. Nogales, E., Wolf, S. G. & Downing, K. H. Structure of the αβ tubulin dimer by electron crystallography. Nature 391, 199–203 (1998).

    CAS  PubMed  Google Scholar 

  163. Elie-Caille, C. et al. Straight GDP-tubulin protofilaments form in the presence of taxol. Curr. Biol. 17, 1765–1770 (2007).

    CAS  PubMed  Google Scholar 

  164. Castle, B. T. et al. Mechanisms of kinetic stabilization by the drugs paclitaxel and vinblastine. Mol. Biol. Cell 28, 1238–1257 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  165. Kellogg, E. H. et al. Insights into the distinct mechanisms of action of taxane and non-taxane microtubule stabilizers from cryo-EM structures. J. Mol. Biol. 429, 633–646 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. Derry, W. B., Wilson, L. & Jordan, M. A. Substoichiometric binding of taxol suppresses microtubule dynamics. Biochemistry 34, 2203–2211 (1995).

    CAS  PubMed  Google Scholar 

  167. Rai, A. et al. Taxanes convert regions of perturbed microtubule growth into rescue sites. Nat. Mater. 19, 355–365 (2020).

    CAS  PubMed  Google Scholar 

  168. Prota, A. E. et al. Structural basis of microtubule stabilization by laulimalide and peloruside A. Angew. Chem. Int. Ed. 53, 1621–1625 (2014).

    CAS  Google Scholar 

  169. McLoughlin, E. C. & O’Boyle, N. M. Colchicine-binding site inhibitors from chemistry to clinic: a review. Pharmaceuticals 13, 8 (2020).

    CAS  PubMed Central  Google Scholar 

  170. Panda, D., Daijo, J. E., Jordan, M. A. & Wilson, L. Kinetic stabilization of microtubule dynamics at steady state in vitro by substoichiometric concentrations of tubulin–colchicine complex. Biochemistry 34, 9921–9929 (1995).

    CAS  PubMed  Google Scholar 

  171. Ranaivoson, F. M., Gigant, B., Berritt, S., Joullié, M. & Knossow, M. Structural plasticity of tubulin assembly probed by vinca-domain ligands. Acta Cryst. D. 68, 927–934 (2012).

    CAS  Google Scholar 

  172. Gigant, B. et al. Structural basis for the regulation of tubulin by vinblastine. Nature 435, 519–522 (2005).

    CAS  PubMed  Google Scholar 

  173. Martino, E. et al. Vinca alkaloids and analogues as anti-cancer agents: looking back, peering ahead. Bioorg. Med. Chem. Lett. 28, 2816–2826 (2018).

    PubMed  Google Scholar 

  174. Mougalian, S. S., Kish, J. K., Zhang, J., Liassou, D. & Feinberg, B. A. Effectiveness of eribulin in metastatic breast cancer: 10 years of real-world clinical experience in the United States. Adv. Ther. 38, 2213–2225 (2021).

    CAS  PubMed  PubMed Central  Google Scholar 

  175. Smith, J. A. et al. Eribulin binds at microtubule ends to a single site on tubulin to suppress dynamic instability. Biochemistry 49, 1331–1337 (2010).

    CAS  PubMed  Google Scholar 

  176. Doodhi, H. et al. Termination of protofilament elongation by eribulin induces lattice defects that promote microtubule catastrophes. Curr. Biol. 26, 1713–1721 (2016).

    CAS  PubMed  Google Scholar 

  177. Prota, A. E. et al. A new tubulin-binding site and pharmacophore for microtubule-destabilizing anticancer drugs. Proc. Natl Acad. Sci. USA 111, 13817–13821 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. Bañuelos-Hernández, A. E., Mendoza-Espinoza, J. A., Pereda-Miranda, R. & Cerda-García-Rojas, C. M. Studies of (−)-pironetin binding to α-tubulin: conformation, docking, and molecular dynamics. J. Org. Chem. 79, 3752–3764 (2014).

    PubMed  Google Scholar 

  179. Chi, Z. & Ambrose, C. Microtubule encounter-based catastrophe in Arabidopsis cortical microtubule arrays. BMC Plant. Biol. 16, 18 (2016).

    PubMed  PubMed Central  Google Scholar 

  180. Laan, L., Husson, J., Munteanu, E. L., Kerssemakers, J. W. J. & Dogterom, M. Force-generation and dynamic instability of microtubule bundles. Proc. Natl Acad. Sci. USA 105, 8920–8925 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  181. Ye, A. A., Cane, S. & Maresca, T. J. Chromosome biorientation produces hundreds of piconewtons at a metazoan kinetochore. Nat. Commun. 7, 13221 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Akiyoshi, B. et al. Tension directly stabilizes reconstituted kinetochore–microtubule attachments. Nature 468, 576–579 (2010). This study discusses microtubule dynamics in vitro and the ways that both a kinetochore component from yeast and applied forces can alter rates of growth and shortening.

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Maiato, H., Gomes, A. M., Sousa, F. & Barisic, M. Mechanisms of chromosome congression during mitosis. Biology 6, 13 (2017).

    PubMed Central  Google Scholar 

  184. Volkov, V. A., Huis in’t Veld, P. J., Dogterom, M. & Musacchio, A. Multivalency of NDC80 in the outer kinetochore is essential to track shortening microtubules and generate forces. eLife 7, e36764 (2018).

    PubMed  PubMed Central  Google Scholar 

  185. Helgeson, L. A. et al. Human Ska complex and Ndc80 complex interact to form a load-bearing assembly that strengthens kinetochore–microtubule attachments. Proc. Natl Acad. Sci. USA 115, 2740–2745 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  186. Huis in’t Veld, P. J., Volkov, V. A., Stender, I. D., Musacchio, A. & Dogterom, M. Molecular determinants of the Ska–Ndc80 interaction and their influence on microtubule tracking and force-coupling. eLife 8, e49539 (2019).

    Google Scholar 

  187. Driver, J. W., Geyer, E. A., Bailey, M. E., Rice, L. M. & Asbury, C. L. Direct measurement of conformational strain energy in protofilaments curling outward from disassembling microtubule tips. eLife 6, e28433 (2017).

    PubMed  PubMed Central  Google Scholar 

  188. Maiato, H., DeLuca, J., Salmon, E. D. & Earnshaw, W. C. The dynamic kinetochore–microtubule interface. J. Cell Sci. 117, 5461–5477 (2004).

    CAS  PubMed  Google Scholar 

  189. Wan, X., Cimini, D., Cameron, L. A. & Salmon, E. D. The coupling between sister kinetochore directional instability and oscillations in centromere stretch in metaphase PtK1 cells. Mol. Biol. Cell 23, 1035–1046 (2012). This thoughtful study investigates the relationship between chromosome motions during prometaphase and the dynamics of microtubules that are required for these motions.

    CAS  PubMed  PubMed Central  Google Scholar 

  190. Trushko, A., Schäffer, E. & Howard, J. The growth speed of microtubules with XMAP215-coated beads coupled to their ends is increased by tensile force. Proc. Natl Acad. Sci. USA 110, 14670–14675 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  191. Volkov, V. A. et al. Long tethers provide high-force coupling of the Dam1 ring to shortening microtubules. Proc. Natl Acad. Sci. USA 110, 7708–7713 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  192. Chakraborti, S., Natarajan, K., Curiel, J., Janke, C. & Liu, J. The emerging role of the tubulin code: from the tubulin molecule to neuronal function and disease. Cytoskeleton 73, 521–550 (2016).

    CAS  PubMed  Google Scholar 

  193. Fees, C. P. & Moore, J. K. Regulation of microtubule dynamic instability by the carboxy-terminal tail of β-tubulin. Life Sci. Alliance 1, e201800054 (2018).

    PubMed  PubMed Central  Google Scholar 

  194. Chen, J. et al. α-Tubulin tail modifications regulate microtubule stability through selective effector recruitment, not changes in intrinsic polymer dynamics. Dev. Cell https://doi.org/10.1016/j.devcel.2021.05.005 (2021).

    Article  PubMed  PubMed Central  Google Scholar 

  195. Young, G. et al. Quantitative mass imaging of single biological macromolecules. Science 360, 423–427 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  196. Nievergelt, A. P., Banterle, N., Andany, S. H., Gönczy, P. & Fantner, G. E. High-speed photothermal off-resonance atomic force microscopy reveals assembly routes of centriolar scaffold protein SAS-6. Nat. Nanotechnol. 13, 696–701 (2018).

    CAS  PubMed  Google Scholar 

  197. Mickolajczyk, K. J., Geyer, E. A., Kim, T., Rice, L. M. & Hancock, W. O. Direct observation of individual tubulin dimers binding to growing microtubules. Proc. Natl Acad. Sci. USA 116, 7314–7322 (2019). This paper presents an interesting use of interferometric scattering microscopy to study the growth of microtubules by the addition of labelled subunits.

    CAS  PubMed  PubMed Central  Google Scholar 

  198. van Haren, J. et al. Local control of intracellular microtubule dynamics by EB1 photodissociation. Nat. Cell Biol. 20, 252–261 (2018).

    PubMed  PubMed Central  Google Scholar 

  199. Aumeier, C. et al. Self-repair promotes microtubule rescue. Nat. Cell Biol. 18, 1054–1064 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  200. Finkenstaedt-Quinn, S. A., Ge, S. & Haynes, C. L. Cytoskeleton dynamics in drug-treated platelets. Anal. Bioanal. Chem. 407, 2803–2809 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  201. Nithianantham, S. et al. Structural basis of tubulin recruitment and assembly by microtubule polymerases with tumor overexpressed gene (TOG) domain arrays. eLife 7, e38922 (2018).

    PubMed  PubMed Central  Google Scholar 

  202. Tran, P. T., Joshi, P. & Salmon, E. D. How tubulin subunits are lost from the shortening ends of microtubules. J. Struct. Biol. 118, 107–118 (1997).

    CAS  PubMed  Google Scholar 

  203. Demchouk, A. O., Gardner, M. K. & Odde, D. J. Microtubule tip tracking and tip structures at the nanometer scale using digital fluorescence microscopy. Cel. Mol. Bioeng. 4, 192–204 (2011).

    Google Scholar 

  204. Kristofferson, D., Mitchison, T. & Kirschner, M. Direct observation of steady-state microtubule dynamics. J. Cell Biol. 102, 1007–1019 (1986).

    CAS  PubMed  Google Scholar 

  205. Snaith, H. A., Anders, A., Samejima, I. & Sawin, K. E. New and old reagents for fluorescent protein tagging of microtubules in fission yeast; experimental and critical evaluation. Methods Cell Biol. 97, 147–172 (2010).

    CAS  PubMed  Google Scholar 

  206. Vala, M. et al. Nanoscopic structural fluctuations of disassembling microtubules revealed by label-free super-resolution microscopy. Small Methods 5, 2000985 (2021).

    CAS  Google Scholar 

  207. Delgado, L., Baeza, N., Pérez-Cruz, C., López-Iglesias, C. & Mercadé, E. Cryo-transmission electron microscopy of outer-inner membrane vesicles naturally secreted by Gram-negative pathogenic bacteria. Bio Protoc. 9, e3367 (2019).

    PubMed  PubMed Central  Google Scholar 

  208. Dehaoui, A., Issenmann, B. & Caupin, F. Viscosity of deeply supercooled water and its coupling to molecular diffusion. Proc. Natl. Acad. Sci. USA 112, 12020–12025 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  209. Grant, T. & Grigorieff, N. Measuring the optimal exposure for single particle cryo-EM using a 2.6 Å reconstruction of rotavirus VP6. eLife 4, e06980 (2015).

    PubMed  PubMed Central  Google Scholar 

  210. Keya, J. J. et al. High-resolution imaging of a single gliding protofilament of tubulins by HS-AFM. Sci. Rep. 7, 6166 (2017).

    PubMed  PubMed Central  Google Scholar 

  211. Dogterom, M. & Yurke, B. Measurement of the force-velocity relation for growing microtubules. Science 278, 856–860 (1997). This paper presents the first measurements on forces developed by microtubules polymerizing in vitro.

    CAS  PubMed  Google Scholar 

  212. Grishchuk, E. L., Molodtsov, M. I., Ataullakhanov, F. I. & McIntosh, J. R. Force production by disassembling microtubules. Nature 438, 384–388 (2005). This paper presents the first measurements on forces developed by microtubules depolymerizing in vitro.

    CAS  PubMed  Google Scholar 

  213. Drechsler, H., Xu, Y., Geyer, V. F., Zhang, Y. & Diez, S. Multivalent electrostatic microtubule interactions of synthetic peptides are sufficient to mimic advanced MAP-like behavior. MBoC 30, 2953–2968 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  214. McIntosh, J. R., Volkov, V., Ataullakhanov, F. I. & Grishchuk, E. L. Tubulin depolymerization may be an ancient biological motor. J. Cell Sci. 123, 3425–3434 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  215. Schmidt, J. C. et al. The kinetochore-bound Ska1 complex tracks depolymerizing microtubules and binds to curved protofilaments. Dev. Cell 23, 968–980 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  216. Grishchuk, E. L. et al. Different assemblies of the DAM1 complex follow shortening microtubules by distinct mechanisms. Proc Natl Acad Sci USA 105, 6918–6923 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  217. Westermann, S. et al. The Dam1 kinetochore ring complex moves processively on depolymerizing microtubule ends. Nature 440, 565–569 (2006).

    CAS  PubMed  Google Scholar 

  218. Miranda, J. L., Wulf, P. D., Sorger, P. K. & Harrison, S. C. The yeast DASH complex forms closed rings on microtubules. Nat. Struct. Mol. Biol. 12, 138 (2005).

    CAS  PubMed  Google Scholar 

  219. Franck, A. D. et al. Tension applied through the Dam1 complex promotes microtubule elongation providing a direct mechanism for length control in mitosis. Nat. Cell Biol. 9, 832–837 (2007). This paper is a direct demonstration of the impact of applied forces on the dynamics of microtubules in vitro.

    CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

The authors thank V. Alexandrova for critical reading of the manuscript and I. Lopanskaia for assistance with the figures. This work was partly supported by National Institutes of Health (NIH) grant GM033787 to J.R.M. Work on microtubule control by regulatory proteins was supported by Russian Foundation for Basic Research grant # 20-34-70159 and work on microtubule control by small-molecule inhibitors was supported by Russian Science Foundation grant # 21-74-20035 to N.B.G.

Author information

Authors and Affiliations

  1. Department of Physics, Lomonosov Moscow State University, Moscow, Russia

    Nikita B. Gudimchuk

  2. Center for Theoretical Problems of Physicochemical Pharmacology, Russian Academy of Sciences, Moscow, Russia

    Nikita B. Gudimchuk

  3. Dmitry Rogachev National Research Center of Pediatric Hematology, Oncology and Immunology, Moscow, Russia

    Nikita B. Gudimchuk

  4. Department of Molecular, Cellular and Developmental Biology, University of Colorado, Boulder, CO, USA

    J. Richard McIntosh

Authors
  1. Nikita B. Gudimchuk
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. J. Richard McIntosh
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

The authors contributed equally to all aspects of the article.

Corresponding authors

Correspondence to Nikita B. Gudimchuk or J. Richard McIntosh.

Ethics declarations

Competing interests

The authors declare that there is no conflict of interest.

Additional information

Peer review information

Nature Reviews Molecular Cell Biology thanks T. Surrey, who co-reviewed with J. Estevez-Gallego, R. Ohi and L. Rice, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Glossary

Ran-GTP

GTP-bound Ran (a member of the Ras superfamily, thus serving as a regulator of biological processes) that stimulates microtubule polymerization by activating several relevant proteins. During mitosis, the concentration of this complex is highest in the vicinity of chromosomes because a chromatin-localized nucleotide exchange factor facilitates conversion of freely diffusing Ran-GDP and GTP into Ran-GTP and GDP.

Formins

Conserved actin polymerases.

Microtubule organizing centre

A cellular structure that nucleates microtubule polymerization in cells.

Focal adhesions

Large protein assemblies that mechanically link the extracellular matrix to cytoplasmic bundles of actin.

Paclitaxel

A small molecule, derived from bark of the Pacific Yew tree Taxus brevifolia, which binds to microtubules and stabilizes them against depolymerization.

Nocodazole

A small molecule that binds soluble tubulin and prevents its polymerization. Nocodazole therefore works as a microtubule-destabilizing agent.

Filopodia

Thin, membrane-coated, actin-rich protrusions of the cell surface.

Immunological synapse

A junction between a T cell and an antigen-presenting cell.

Microtubule cross-linkers

Proteins that connect two or more microtubules into a bundle.

Combrestatin

A small-molecule microtubule-destabilizing agent derived from the African plant Combretum caffrum.

GMPCPP

A slowly hydrolysable analogue of GTP in which the α-phosphates and β-phosphates are connected via a methylene link.

Colchicine

A small molecule derived from the plant Colchicum autumnale. It binds tubulin and works as a microtubule-destabilizing agent.

Microtubule-associated proteins

A heterogeneous group of proteins that bind to microtubules. The group includes motor enzymes, some microtubule dynamics regulators, microtubule cross-linkers and so on.

Axoneme

A structure comprising the core of a eukaryotic cilium or flagellum. In primary (non-motile) cilia, it consists of nine microtubule doublets arranged in a cylinder. Motile cilia and flagella additionally have a pair of microtubules in the centre of the cylinder. Microtubules within the axoneme are bridged by multiple cross-linking proteins, including dynein motor proteins, which enable sliding of adjacent microtubule doublets to produce a beating motion.

TOG domain-containing proteins

Microtubule-associated proteins that contain from two to five TOG (Tumour Overexpressed Gene) domains. These are tubulin-binding domains that can affect microtubule dynamics.

Epothilone

A small molecule, derived from the myxobacterium Sorangium cellulosum, which acts as a microtubule-stabilizing agent.

Kinetochores

Large protein assemblies that form at a chromosome’s centromere. They are responsible for microtubule capture in mitosis.

Microtubule flux

A cellular process in which a microtubule moves along its axis towards its minus end while tubulin subunits are added at its plus end and removed from its minus end. The microtubule moves, but both its ends and its centre of mass are stationary.

Anaphase B

The second part of anaphase. During this stage of cell division, the mitotic spindle elongates through the growth and sliding apart of antiparallel, interpolar microtubules.

Growth cones

Motile, actin-rich cellular specializations at the tips of neuronal extensions, such as axons and dendrites. Their motility draws the tips of a developing neuronal branch and enables the formation of neural connections.

Dolastatin

A collective name for several microtubule-inhibiting peptides, derived from a mollusc from the Indian Ocean, Dolabella auricularia. The peptides bind tubulin and promote the formation of curved protofilament-like structure that commonly assemble into rings.

Peloruside A

A small-molecule microtubule-stabilizing agent isolated from the New Zealand marine sponge Mycale hentscheli.

Laulimalide

A small-molecule microtubule-stabilizing agent isolated from the marine sponge Cacospongia mycofijiensis.

Vinca-domain ligands

Small molecules that destabilize microtubule assembly through binding to the same site on tubulin as vinblastine, an alkaloid derived from the Madagascar periwinkle plant Vinca rosea.

Eribulin

A small-molecule microtubule-destabilizing agent. It is a synthetic analogue of a polyether macrolide derived from the marine sponge Halichondria okadai.

Maytansine

A small-molecule microtubule-destabilizing agent derived from the plant Maytenus ovatus.

Pironetin

A small-molecule microtubule-destabilizing agent derived from the bacterium Streptomyces prunicolor.

‘Brownian ratchet’ mechanism

Any process in which directed motion of a small particle is achieved through ‘rectification’ of thermal fluctuations, fuelled by some external energy source, such as the growth of a polymer.

Atomic force microscopy

A high-resolution, non-optical imaging method in which structural information about the sample is collected by raster-scanning a very thin, cylindrical probe with a sharp tip over a sample’s surface, using an optical feedback loop to adjust the parameters needed for imaging.

Interferometric scattering microscopy

A sensitive method for optical microscopy in which contrast is achieved through interference of light scattered by the imaged particle with a reference wave that is partially reflected by the microscope slide or coverslip.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Gudimchuk, N.B., McIntosh, J.R. Regulation of microtubule dynamics, mechanics and function through the growing tip. Nat Rev Mol Cell Biol 22, 777–795 (2021). https://doi.org/10.1038/s41580-021-00399-x

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41580-021-00399-x

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing