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.
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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.
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).
Goodson, H. V. & Jonasson, E. M. Microtubules and microtubule-associated proteins. Cold Spring Harb. Perspect. Biol. 10, a022608 (2018).
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).
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).
Kanfer, G. et al. CENP-F couples cargo to growing and shortening microtubule ends. Mol. Biol. Cell 28, 2400–2409 (2017).
Kirschner, M. W. & Mitchison, T. Microtubule dynamics. Nature 324, 621 (1986).
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).
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).
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).
Dogterom, M. & Koenderink, G. H. Actin–microtubule crosstalk in cell biology. Nat. Rev. Mol. Cell Biol. 20, 38–54 (2019).
Seetharaman, S. & Etienne-Manneville, S. Cytoskeletal crosstalk in cell migration. Trends Cell Biol. 30, 720–735 (2020).
Jiang, K. et al. A proteome-wide screen for mammalian SxIP motif-containing microtubule plus-end tracking proteins. Curr. Biol. 22, 1800–1807 (2012).
Kodama, A., Karakesisoglou, I., Wong, E., Vaezi, A. & Fuchs, E. ACF7: an essential integrator of microtubule dynamics. Cell 115, 343–354 (2003).
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).
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).
Daga, R. R., Yonetani, A. & Chang, F. Asymmetric microtubule pushing forces in nuclear centering. Curr. Biol. 16, 1544–1550 (2006).
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).
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).
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).
Penfield, L. et al. Dynein pulling forces counteract lamin-mediated nuclear stability during nuclear envelope repair. MBoC 29, 852–868 (2018).
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).
Tanimoto, H., Kimura, A. & Minc, N. Shape–motion relationships of centering microtubule asters. J. Cell Biol. 212, 777–787 (2016).
Laan, L. et al. Cortical dynein controls microtubule dynamics to generate pulling forces that position microtubule asters. Cell 148, 502–514 (2012).
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).
Burakov, A., Nadezhdina, E., Slepchenko, B. & Rodionov, V. Centrosome positioning in interphase cells. J. Cell Biol. 162, 963–969 (2003).
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).
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).
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).
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).
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.
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.
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).
Vukušic´, K., Buda, R. & Tolic´, I. M. Force-generating mechanisms of anaphase in human cells. J. Cell Sci. 132, jcs231985 (2019).
McIntosh, J. R. Anaphase A. Semin. Cell Dev. Biol. https://doi.org/10.1016/j.semcdb.2021.03.009 (2021).
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).
Heald, R. & Khodjakov, A. Thirty years of search and capture: the complex simplicity of mitotic spindle assembly. J. Cell Biol. 211, 1103–1111 (2015).
Röper, K. Microtubules enter centre stage for morphogenesis. Philos. Trans. R. Soc. B https://doi.org/10.1098/rstb.2019.0557 (2020).
Kaverina, I., Rottner, K. & Small, J. V. Targeting, capture, and stabilization of microtubules at early focal adhesions. J. Cell Biol. 142, 181–190 (1998).
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).
Kopf, A. et al. Microtubules control cellular shape and coherence in amoeboid migrating cells. J. Cell Biol. 219, e201907154 (2020).
Tabdanov, E. D. et al. Engineering T cells to enhance 3D migration through structurally and mechanically complex tumor microenvironments. Nat. Commun. 12, 2815 (2021).
Schelski, M. & Bradke, F. Neuronal polarization: from spatiotemporal signaling to cytoskeletal dynamics. Mol. Cell. Neurosci. 84, 11–28 (2017).
Witte, H., Neukirchen, D. & Bradke, F. Microtubule stabilization specifies initial neuronal polarization. J. Cell Biol. 180, 619–632 (2008).
Dupraz, S. et al. RhoA controls axon extension independent of specification in the developing brain. Curr. Biol. 29, 3874–3886.e9 (2019).
Singh, A. et al. Polarized microtubule dynamics directs cell mechanics and coordinates forces during epithelial morphogenesis. Nat. Cell Biol. 20, 1126–1133 (2018).
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).
Diagouraga, B. et al. Motor-driven marginal band coiling promotes cell shape change during platelet activation. J. Cell Biol. 204, 177–185 (2014).
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).
Hooikaas, P. J. et al. Kinesin-4 KIF21B limits microtubule growth to allow rapid centrosome polarization in T cells. eLife 9, e62876 (2020).
McIntosh, J. R., Grishchuk, E. L. & West, R. R. Chromosome–microtubule interactions during mitosis. Annu. Rev. Cell Dev. Biol. 18, 193–219 (2002).
Kapoor, T. M. Metaphase spindle assembly. Biology 6, 8 (2017).
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).
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).
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).
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).
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).
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.
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).
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).
Roostalu, J. et al. The speed of GTP hydrolysis determines GTP cap size and controls microtubule stability. eLife 9, e51992 (2020).
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.
Marantz, R. & Shelanski, M. L. Structure of microtubular crystals induced by vinblastine in vitro. J. Cell Biol. 44, 234–238 (1970).
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).
Wang, H.-W. & Nogales, E. Nucleotide-dependent bending flexibility of tubulin regulates microtubule assembly. Nature 435, 911–915 (2005).
Howes, S. C. et al. Structural and functional differences between porcine brain and budding yeast microtubules. Cell Cycle 17, 278–287 (2018).
Ayukawa, R. et al. GTP-dependent formation of straight tubulin oligomers leads to microtubule nucleation. J. Cell Biol. 220, e202007033 (2021).
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.
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).
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).
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).
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).
Brouhard, G. J. & Rice, L. M. The contribution of αβ-tubulin curvature to microtubule dynamics. J. Cell Biol. 207, 323–334 (2014).
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).
Grafmüller, A. & Voth, G. A. Intrinsic bending of microtubule protofilaments. Structure 19, 409–417 (2011).
Igaev, M. & Grubmüller, H. Microtubule assembly governed by tubulin allosteric gain in flexibility and lattice induced fit. eLife 7, e34353 (2018).
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).
Tong, D. & Voth, G. A. Microtubule simulations provide insight into the molecular mechanism underlying dynamic instability. Biophys. J. 118, 2938–2951 (2020).
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.
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.
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).
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.
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).
Howes, S. C. et al. Structural differences between yeast and mammalian microtubules revealed by cryo-EM. J. Cell Biol. 216, 2669–2677 (2017).
Estévez-Gallego, J. et al. Structural model for differential cap maturation at growing microtubule ends. eLife 9, e50155 (2020).
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.
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).
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).
Zakharov, P. et al. Molecular and mechanical causes of microtubule catastrophe and aging. Biophys. J. 109, 2574–2591 (2015).
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.
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).
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).
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).
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.
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.
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).
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).
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).
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).
VandenBeldt, K. J. et al. Kinetochores use a novel mechanism for coordinating the dynamics of individual microtubules. Curr. Biol. 16, 1217–1223 (2006).
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).
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).
Hoog, J. L. et al. Electron tomography reveals a flared morphology on growing microtubule ends. J. Cell Sci. 124, 693–698 (2011).
Kukulski, W. et al. Correlated fluorescence and 3D electron microscopy with high sensitivity and spatial precision. J. Cell Biol. 192, 111–119 (2011).
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).
Vitre, B. et al. EB1 regulates microtubule dynamics and tubulin sheet closure in vitro. Nat. Cell Biol. 10, 415–421 (2008).
Komarova, Y. et al. Mammalian end binding proteins control persistent microtubule growth. J. Cell Biol. 184, 691–706 (2009).
Bieling, P. et al. Reconstitution of a microtubule plus-end tracking system in vitro. Nature 450, 1100–1105 (2007).
Maurer, S. P. et al. EB1 accelerates two conformational transitions important for microtubule maturation and dynamics. Curr. Biol. 24, 372–384 (2014).
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).
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).
Gardner, M. K. et al. Rapid microtubule self-assembly kinetics. Cell 146, 582–592 (2011).
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).
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).
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).
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).
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).
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).
Miller, M. P., Asbury, C. L. & Biggins, S. A TOG protein confers tension sensitivity to kinetochore–microtubule attachments. Cell 165, 1428 (2016).
Al-Bassam, J. & Chang, F. Regulation of microtubule dynamics by TOG-domain proteins XMAP215/Dis1 and CLASP. Trends Cell Biol. 21, 604–614 (2011).
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).
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.
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).
Byrnes, A. E. & Slep, K. C. TOG–tubulin binding specificity promotes microtubule dynamics and mitotic spindle formation. J. Cell Biol. 216, 1641–1657 (2017).
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).
Brouhard, G. J. et al. XMAP215 is a processive microtubule polymerase. Cell 132, 79–88 (2008).
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).
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).
Ayaz, P. et al. A tethered delivery mechanism explains the catalytic action of a microtubule polymerase. eLife 3, e03069 (2014).
Drabek, K. et al. Role of CLASP2 in microtubule stabilization and the regulation of persistent motility. Curr. Biol. 16, 2259–2264 (2006).
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).
Majumdar, S. et al. An isolated CLASP TOG domain suppresses microtubule catastrophe and promotes rescue. MBoC 29, 1359–1375 (2018).
Lawrence, E. J., Arpag˘, G., Norris, S. R. & Zanic, M. Human CLASP2 specifically regulates microtubule catastrophe and rescue. MBoC 29, 1168–1177 (2018).
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).
Leano, J. B. & Slep, K. C. Structures of TOG1 and TOG2 from the human microtubule dynamics regulator CLASP1. PLoS ONE 14, e0219823 (2019).
Aher, A. et al. CLASP suppresses microtubule catastrophes through a single TOG domain. Dev. Cell 46, 40–58.e8 (2018).
Aher, A. et al. CLASP mediates microtubule repair by restricting lattice damage and regulating tubulin incorporation. Curr. Biol. 30, 2175–2183.e6 (2020).
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).
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).
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).
Ganem, N. J., Godinho, S. A. & Pellman, D. A mechanism linking extra centrosomes to chromosomal instability. Nature 460, 278 (2009).
Ohi, R., Burbank, K., Liu, Q. & Mitchison, T. J. Nonredundant functions of kinesin-13s during meiotic spindle assembly. Curr. Biol. 17, 953–959 (2007).
Homma, N. et al. Kinesin superfamily protein 2A (KIF2A) functions in suppression of collateral branch extension. Cell 114, 229–239 (2003).
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).
Wang, W. et al. Insight into microtubule disassembly by kinesin-13s from the structure of Kif2C bound to tubulin. Nat. Commun. 8, 70 (2017).
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).
Paydar, M. & Kwok, B. H. Evidence for conformational change-induced hydrolysis of β-tubulin-GTP. Sneak Peak https://papers.ssrn.com/abstract=3687033 (2020)
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).
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).
Varga, V. et al. Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner. Nat. Cell Biol. 8, 957–962 (2006).
Arellano-Santoyo, H. et al. A tubulin binding switch underlies Kip3/Kinesin-8 depolymerase activity. Dev. Cell 42, 37–51.e8 (2017).
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).
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).
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).
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).
Gupta, K. K. et al. Mechanism for the catastrophe-promoting activity of the microtubule destabilizer Op18/stathmin. PNAS 110, 20449–20454 (2013).
Jordan, M. A. & Wilson, L. Microtubules as a target for anticancer drugs. Nat. Rev. Cancer 4, 253–265 (2004).
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).
Steinmetz, M. O. & Prota, A. E. Microtubule-targeting agents: strategies to hijack the cytoskeleton. Trends Cell Biol. 28, 776–792 (2018).
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).
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).
Nogales, E., Wolf, S. G. & Downing, K. H. Structure of the αβ tubulin dimer by electron crystallography. Nature 391, 199–203 (1998).
Elie-Caille, C. et al. Straight GDP-tubulin protofilaments form in the presence of taxol. Curr. Biol. 17, 1765–1770 (2007).
Castle, B. T. et al. Mechanisms of kinetic stabilization by the drugs paclitaxel and vinblastine. Mol. Biol. Cell 28, 1238–1257 (2017).
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).
Derry, W. B., Wilson, L. & Jordan, M. A. Substoichiometric binding of taxol suppresses microtubule dynamics. Biochemistry 34, 2203–2211 (1995).
Rai, A. et al. Taxanes convert regions of perturbed microtubule growth into rescue sites. Nat. Mater. 19, 355–365 (2020).
Prota, A. E. et al. Structural basis of microtubule stabilization by laulimalide and peloruside A. Angew. Chem. Int. Ed. 53, 1621–1625 (2014).
McLoughlin, E. C. & O’Boyle, N. M. Colchicine-binding site inhibitors from chemistry to clinic: a review. Pharmaceuticals 13, 8 (2020).
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).
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).
Gigant, B. et al. Structural basis for the regulation of tubulin by vinblastine. Nature 435, 519–522 (2005).
Martino, E. et al. Vinca alkaloids and analogues as anti-cancer agents: looking back, peering ahead. Bioorg. Med. Chem. Lett. 28, 2816–2826 (2018).
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).
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).
Doodhi, H. et al. Termination of protofilament elongation by eribulin induces lattice defects that promote microtubule catastrophes. Curr. Biol. 26, 1713–1721 (2016).
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).
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).
Chi, Z. & Ambrose, C. Microtubule encounter-based catastrophe in Arabidopsis cortical microtubule arrays. BMC Plant. Biol. 16, 18 (2016).
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).
Ye, A. A., Cane, S. & Maresca, T. J. Chromosome biorientation produces hundreds of piconewtons at a metazoan kinetochore. Nat. Commun. 7, 13221 (2016).
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.
Maiato, H., Gomes, A. M., Sousa, F. & Barisic, M. Mechanisms of chromosome congression during mitosis. Biology 6, 13 (2017).
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).
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).
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).
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).
Maiato, H., DeLuca, J., Salmon, E. D. & Earnshaw, W. C. The dynamic kinetochore–microtubule interface. J. Cell Sci. 117, 5461–5477 (2004).
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.
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).
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).
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).
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).
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).
Young, G. et al. Quantitative mass imaging of single biological macromolecules. Science 360, 423–427 (2018).
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).
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.
van Haren, J. et al. Local control of intracellular microtubule dynamics by EB1 photodissociation. Nat. Cell Biol. 20, 252–261 (2018).
Aumeier, C. et al. Self-repair promotes microtubule rescue. Nat. Cell Biol. 18, 1054–1064 (2016).
Finkenstaedt-Quinn, S. A., Ge, S. & Haynes, C. L. Cytoskeleton dynamics in drug-treated platelets. Anal. Bioanal. Chem. 407, 2803–2809 (2015).
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).
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).
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).
Kristofferson, D., Mitchison, T. & Kirschner, M. Direct observation of steady-state microtubule dynamics. J. Cell Biol. 102, 1007–1019 (1986).
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).
Vala, M. et al. Nanoscopic structural fluctuations of disassembling microtubules revealed by label-free super-resolution microscopy. Small Methods 5, 2000985 (2021).
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).
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).
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).
Keya, J. J. et al. High-resolution imaging of a single gliding protofilament of tubulins by HS-AFM. Sci. Rep. 7, 6166 (2017).
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.
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.
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).
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).
Schmidt, J. C. et al. The kinetochore-bound Ska1 complex tracks depolymerizing microtubules and binds to curved protofilaments. Dev. Cell 23, 968–980 (2012).
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).
Westermann, S. et al. The Dam1 kinetochore ring complex moves processively on depolymerizing microtubule ends. Nature 440, 565–569 (2006).
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).
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.
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.
The authors declare that there is no conflict of interest.
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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.
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.
A small molecule, derived from bark of the Pacific Yew tree Taxus brevifolia, which binds to microtubules and stabilizes them against depolymerization.
A small molecule that binds soluble tubulin and prevents its polymerization. Nocodazole therefore works as a microtubule-destabilizing agent.
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.
A small-molecule microtubule-destabilizing agent derived from the African plant Combretum caffrum.
A slowly hydrolysable analogue of GTP in which the α-phosphates and β-phosphates are connected via a methylene link.
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.
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.
A small molecule, derived from the myxobacterium Sorangium cellulosum, which acts as a microtubule-stabilizing agent.
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.
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.
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.
A small-molecule microtubule-destabilizing agent. It is a synthetic analogue of a polyether macrolide derived from the marine sponge Halichondria okadai.
A small-molecule microtubule-destabilizing agent derived from the plant Maytenus ovatus.
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.
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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
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