The Tubulin Code: A Navigation System for Chromosomes during Mitosis

Trends in Cell Biology

Volumn 26, Issue 10, 2016, Pages 766-775

  Barisic, Marin ^a   Maiato, Helder ^b,c,d  

^a DANISH CANCER SOCIETY RESEARCH CENTER   (Denmark)

^b UNIVERSITY OF PORTO   (Portugal)

^c UNIVERSITY OF PORTO   (Portugal)

^d University of Porto   (Portugal)

Author keywords

CENP E; detyrosination; Dynein; motors; post translational modifications; tubulin code

Indexed keywords

TUBULIN;

CENTROMERE; CHROMOSOME; CHROMOSOME SEGREGATION; HUMAN; MITOSIS; MITOSIS SPINDLE; PRIORITY JOURNAL; PROTEIN PROCESSING; REVIEW; TUBULIN CODE; ANIMAL; KINETOCHORE; METABOLISM; SPINDLE APPARATUS;

ANIMALS; CHROMOSOMES; HUMANS; KINETOCHORES; MITOSIS; PROTEIN PROCESSING, POST-TRANSLATIONAL; SPINDLE APPARATUS; TUBULIN;

EID: 84991794896     PISSN: 09628924     EISSN: 18793088     Source Type: Journal    
DOI: 10.1016/j.tcb.2016.06.001     Document Type: Review

References (98)

1. 1 Schweizer, N., et al. An organelle-exclusion envelope assists mitosis and underlies distinct molecular crowding in the spindle region. J. Cell. Biol. 210 (2015), 695–704.
2. 2 Barisic, M., et al. Kinetochore motors drive congression of peripheral polar chromosomes by overcoming random arm-ejection forces. Nat. Cell Biol. 16 (2014), 1249–1256.
3. 3 Bancroft, J., et al. Chromosome congression is promoted by CENP-Q- and CENP-E-dependent pathways. J. Cell Sci. 128 (2015), 171–184.
4. 4 Magidson, V., et al. The spatial arrangement of chromosomes during prometaphase facilitates spindle assembly. Cell 146 (2011), 555–567.
5. 5 Magidson, V., et al. Adaptive changes in the kinetochore architecture facilitate proper spindle assembly. Nat. Cell Biol. 17 (2015), 1134–1144.
6. 6 Mitchison, T., et al. Sites of microtubule assembly and disassembly in the mitotic spindle. Cell 45 (1986), 515–527.
7. 7 McIntosh, J.R., et al. Tubulin depolymerization may be an ancient biological motor. J. Cell Sci. 123 (2010), 3425–3434.
8. 8 Auckland, P., McAinsh, A.D., Building an integrated model of chromosome congression. J. Cell Sci. 128 (2015), 3363–3374.
9. 9 Li, Y., et al. Kinetochore dynein generates a poleward pulling force to facilitate congression and full chromosome alignment. Cell Res. 17 (2007), 701–712.
10. 10 Rieder, C.L., Alexander, S.P., Kinetochores are transported poleward along a single astral microtubule during chromosome attachment to the spindle in newt lung cells. J. Cell Biol. 110 (1990), 81–95.
11. 11 Vorozhko, V.V., et al. Multiple mechanisms of chromosome movement in vertebrate cells mediated through the Ndc80 complex and dynein/dynactin. Chromosoma 117 (2008), 169–179.
12. 12 Yang, Z., et al. Kinetochore dynein is required for chromosome motion and congression independent of the spindle checkpoint. Curr. Biol. 17 (2007), 973–980.
13. 13 Kapoor, T.M., et al. Chromosomes can congress to the metaphase plate before biorientation. Science 311 (2006), 388–391.
14. 14 Schaar, B.T., et al. CENP-E function at kinetochores is essential for chromosome alignment. J. Cell Biol. 139 (1997), 1373–1382.
15. 15 Wood, K.W., et al. CENP-E is a plus end-directed kinetochore motor required for metaphase chromosome alignment. Cell 91 (1997), 357–366.
16. 16 Antonio, C., et al. Xkid, a chromokinesin required for chromosome alignment on the metaphase plate. Cell 102 (2000), 425–435.
17. 17 Funabiki, H., Murray, A.W., The Xenopus chromokinesin Xkid is essential for metaphase chromosome alignment and must be degraded to allow anaphase chromosome movement. Cell 102 (2000), 411–424.
18. 18 Wandke, C., et al. Human chromokinesins promote chromosome congression and spindle microtubule dynamics during mitosis. J. Cell Biol. 198 (2012), 847–863.
19. 19 Iemura, K., Tanaka, K., Chromokinesin Kid and kinetochore kinesin CENP-E differentially support chromosome congression without end-on attachment to microtubules. Nat. Commun., 6, 2015, 6447.
20. 20 Cane, S., et al. Elevated polar ejection forces stabilize kinetochore-microtubule attachments. J. Cell Biol. 200 (2013), 203–218.
21. 21 Drpic, D., et al. Polar ejection forces promote the conversion from lateral to end-on kinetochore-microtubule attachments on mono-oriented chromosomes. Cell Rep. 13 (2015), 460–469.
22. 22 Lampson, M.A., Cheeseman, I.M., Sensing centromere tension: Aurora B and the regulation of kinetochore function. Trends Cell Biol. 21 (2011), 133–140.
23. 23 Chmatal, L., et al. Spatial regulation of kinetochore microtubule attachments by destabilization at spindle poles in meiosis I. Curr. Biol. 25 (2015), 1835–1841.
24. 24 Ye, A.A., et al. Aurora A kinase contributes to a pole-based error correction pathway. Curr. Biol. 25 (2015), 1842–1851.
25. 25 Barisic, M., Maiato, H., Dynein prevents erroneous kinetochore-microtubule attachments in mitosis. Cell Cycle 14 (2015), 3356–3361.
26. 26 Kim, Y., et al. Aurora kinases and protein phosphatase 1 mediate chromosome congression through regulation of CENP-E. Cell 142 (2010), 444–455.
27. 27 Cai, S., et al. Chromosome congression in the absence of kinetochore fibres. Nat. Cell Biol. 11 (2009), 832–838.
28. 28 Hasegawa, K., et al. Chromosomal gain promotes formation of a steep RanGTP gradient that drives mitosis in aneuploid cells. J. Cell Biol. 200 (2013), 151–161.
29. 29 Barisic, M., et al. Mitosis. Microtubule detyrosination guides chromosomes during mitosis. Science (New York, N.Y.). 348 (2015), 799–803.
30. 30 Jenuwein, T., Allis, C.D., Translating the histone code. Science 293 (2001), 1074–1080.
31. 31 Janke, C., The tubulin code: molecular components, readout mechanisms, and functions. J. Cell Biol. 206 (2014), 461–472.
32. 32 Verhey, K.J., Gaertig, J., The tubulin code. Cell Cycle 6 (2007), 2152–2160.
33. 33 Fourest-Lieuvin, A., et al. Microtubule regulation in mitosis: tubulin phosphorylation by the cyclin-dependent kinase Cdk1. Mol. Biol. Cell 17 (2006), 1041–1050.
34. 34 Caudron, F., et al. Mutation of Ser172 in yeast beta tubulin induces defects in microtubule dynamics and cell division. PLoS One, 5, 2010, e13553.
35. 35 Hallak, M.E., et al. Release of tyrosine from tyrosinated tubulin Some common factors that affect this process and the assembly of tubulin. FEBS Lett. 73 (1977), 147–150.
36. 36 Raybin, D., Flavin, M., An enzyme tyrosylating alpha-tubulin and its role in microtubule assembly. Biochem. Biophys. Res. Commun. 65 (1975), 1088–1095.
37. 37 Schroder, H.C., et al. Purification of brain tubulin-tyrosine ligase by biochemical and immunological methods. J. Cell Biol. 100 (1985), 276–281.
38. 38 Ersfeld, K., et al. Characterization of the tubulin-tyrosine ligase. J. Cell Biol. 120 (1993), 725–732.
39. 39 Akella, J.S., et al. MEC-17 is an alpha-tubulin acetyltransferase. Nature 467 (2010), 218–222.
40. 40 Shida, T., et al. The major alpha-tubulin K40 acetyltransferase alphaTAT1 promotes rapid ciliogenesis and efficient mechanosensation. Proc. Natl. Acad. Sci. U.S.A. 107 (2010), 21517–21522.
41. 41 Hubbert, C., et al. HDAC6 is a microtubule-associated deacetylase. Nature 417 (2002), 455–458.
42. 42 North, B.J., et al. The human Sir2 ortholog, SIRT2, is an NAD+-dependent tubulin deacetylase. Mol. Cell 11 (2003), 437–444.
43. 43 Regnard, C., et al. Tubulin polyglutamylase: partial purification and enzymatic properties. Biochemistry 37 (1998), 8395–8404.
44. 44 Janke, C., et al. Tubulin polyglutamylase enzymes are members of the TTL domain protein family. Science (New York, N.Y.). 308 (2005), 1758–1762.
45. 45 van Dijk, J., et al. A targeted multienzyme mechanism for selective microtubule polyglutamylation. Mol. Cell 26 (2007), 437–448.
46. 46 Ikegami, K., et al. TTLL7 is a mammalian beta-tubulin polyglutamylase required for growth of MAP2-positive neurites. J. Biol. Chem. 281 (2006), 30707–30716.
47. 47 Kimura, Y., et al. Identification of tubulin deglutamylase among Caenorhabditis elegans and mammalian cytosolic carboxypeptidases (CCPs). J. Biol. Chem. 285 (2010), 22936–22941.
48. 48 Rogowski, K., et al. A family of protein-deglutamylating enzymes associated with neurodegeneration. Cell 143 (2010), 564–578.
49. 49 Tort, O., et al. The cytosolic carboxypeptidases CCP2 and CCP3 catalyze posttranslational removal of acidic amino acids. Mol. Biol. Cell 25 (2014), 3017–3027.
50. 50 Bobinnec, Y., et al. Glutamylation of centriole and cytoplasmic tubulin in proliferating non-neuronal cells. Cell Motil. Cytoskeleton 39 (1998), 223–232.
51. 51 Gundersen, G.G., Bulinski, J.C., Distribution of tyrosinated and nontyrosinated alpha-tubulin during mitosis. J. Cell Biol. 102 (1986), 1118–1126.
52. 52 Gundersen, G.G., et al. Distinct populations of microtubules: tyrosinated and nontyrosinated alpha tubulin are distributed differently in vivo. Cell 38 (1984), 779–789.
53. 53 Wilson, P.J., Forer, A., Effects of nanomolar taxol on crane-fly spermatocyte spindles indicate that acetylation of kinetochore microtubules can be used as a marker of poleward tubulin flux. Cell Motil. Cytoskeleton 37 (1997), 20–32.
54. 54 Khawaja, S., et al. Enhanced stability of microtubules enriched in detyrosinated tubulin is not a direct function of detyrosination level. J. Cell Biol. 106 (1988), 141–149.
55. 55 Webster, D.R., et al. Detyrosination of alpha tubulin does not stabilize microtubules in vivo. J. Cell Biol. 111 (1990), 113–122.
56. 56 Maruta, H., et al. The acetylation of alpha-tubulin and its relationship to the assembly and disassembly of microtubules. J. Cell Biol. 103 (1986), 571–579.
57. 57 Paturle, L., et al. Complete separation of tyrosinated, detyrosinated, and nontyrosinatable brain tubulin subpopulations using affinity chromatography. Biochemistry 28 (1989), 2698–2704.
58. 58 Piperno, G., et al. Microtubules containing acetylated alpha-tubulin in mammalian cells in culture. J. Cell Biol. 104 (1987), 289–302.
59. 59 Hammond, J.W., et al. Posttranslational modifications of tubulin and the polarized transport of kinesin-1 in neurons. Mol. Biol. Cell 21 (2010), 572–583.
60. 60 Kalebic, N., et al. Tubulin acetyltransferase alphaTAT1 destabilizes microtubules independently of its acetylation activity. Mol Cell Biol 33 (2013), 1114–1123.
61. 61 Zilberman, Y., et al. Regulation of microtubule dynamics by inhibition of the tubulin deacetylase HDAC6. J. Cell Sci. 122 (2009), 3531–3541.
62. 62 Peris, L., et al. Motor-dependent microtubule disassembly driven by tubulin tyrosination. J. Cell Biol. 185 (2009), 1159–1166.
63. 63 Sirajuddin, M., et al. Regulation of microtubule motors by tubulin isotypes and post-translational modifications. Nat. Cell Biol. 16 (2014), 335–344.
64. 64 Lacroix, B., et al. Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. J. Cell Biol. 189 (2010), 945–954.
65. 65 Valenstein, M.L., Roll-Mecak, A., Graded control of microtubule severing by tubulin glutamylation. Cell 164 (2016), 911–921.
66. 66 Konishi, Y., Setou, M., Tubulin tyrosination navigates the kinesin-1 motor domain to axons. Nat. Neurosci. 12 (2009), 559–567.
67. 67 Maas, C., et al. Synaptic activation modifies microtubules underlying transport of postsynaptic cargo. Proc. Natl. Acad. Sci. U.S.A. 106 (2009), 8731–8736.
68. 68 Reed, N.A., et al. Microtubule acetylation promotes kinesin-1 binding and transport. Curr. Biol. 16 (2006), 2166–2172.
69. 69 Kaul, N., et al. Effects of alpha-tubulin K40 acetylation and detyrosination on kinesin-1 motility in a purified system. Biophys. J. 106 (2014), 2636–2643.
70. 70 Soppina, V., et al. Luminal localization of alpha-tubulin K40 acetylation by cryo-EM analysis of fab-labeled microtubules. PLoS One, 7, 2012, e48204.
71. 71 Szyk, A., et al. Molecular basis for age-dependent microtubule acetylation by tubulin acetyltransferase. Cell 157 (2014), 1405–1415.
72. 72 Whipple, R.A., et al. Parthenolide and costunolide reduce microtentacles and tumor cell attachment by selectively targeting detyrosinated tubulin independent from NF-kappaB inhibition. Breast Cancer Res., 15, 2013, R83.
73. 73 Fonrose, X., et al. Parthenolide inhibits tubulin carboxypeptidase activity. Cancer Res. 67 (2007), 3371–3378.
74. 74 Wang, Z., Sheetz, M.P., The C-terminus of tubulin increases cytoplasmic dynein and kinesin processivity. Biophys. J. 78 (2000), 1955–1964.
75. 75 McKenney, R.J., et al. Activation of cytoplasmic dynein motility by dynactin-cargo adapter complexes. Science (New York, N.Y.). 345 (2014), 337–341.
76. 76 Alper, J.D., et al. The motility of axonemal dynein is regulated by the tubulin code. Biophys. J. 107 (2014), 2872–2880.
77. 77 Kubo, T., et al. Tubulin polyglutamylation regulates axonemal motility by modulating activities of inner-arm dyneins. Curr. Biol. 20 (2010), 441–445.
78. 78 Peris, L., et al. Tubulin tyrosination is a major factor affecting the recruitment of CAP-Gly proteins at microtubule plus ends. J. Cell Biol. 174 (2006), 839–849.
79. 79 Eiserich, J.P., et al. Microtubule dysfunction by posttranslational nitrotyrosination of alpha-tubulin: a nitric oxide-dependent mechanism of cellular injury. Proc. Natl. Acad. Sci. U.S.A. 96 (1999), 6365–6370.
80. 80 McKenney, R.J., et al. Tyrosination of alpha-tubulin controls the initiation of processive dynein-dynactin motility. EMBO J., 2016.
81. 81 Nirschl, J.J., et al. alpha-Tubulin tyrosination and CLIP-170 phosphorylation regulate the initiation of dynein-driven transport in neurons. Cell Rep. 14 (2016), 2637–2652.
82. 82 Erck, C., et al. A vital role of tubulin-tyrosine-ligase for neuronal organization. Proc. Natl. Acad. Sci. U.S.A. 102 (2005), 7853–7858.
83. 83 Kalebic, N., et al. alphaTAT1 is the major alpha-tubulin acetyltransferase in mice. Nat. Commun., 4, 2013, 1962.
84. 84 Bosch Grau, M., et al. Tubulin glycylases and glutamylases have distinct functions in stabilization and motility of ependymal cilia. J. Cell Biol. 202 (2013), 441–451.
85. 85 Rocha, C., et al. Tubulin glycylases are required for primary cilia, control of cell proliferation and tumor development in colon. EMBO J. 33 (2014), 2247–2260.
86. 86 Vemu, A., et al. Generation of differentially modified microtubules using in vitro enzymatic approaches. Methods Enzymol. 540 (2014), 149–166.
87. 87 Liu, N., et al. Proteomic profiling and functional characterization of multiple post-translational modifications of tubulin. J. Proteome Res. 14 (2015), 3292–3304.
88. 88 Lafanechere, L., et al. Suppression of tubulin tyrosine ligase during tumor growth. J. Cell Sci. 111:Pt 2 (1998), 171–181.
89. 89 Mialhe, A., et al. Tubulin detyrosination is a frequent occurrence in breast cancers of poor prognosis. Cancer Res. 61 (2001), 5024–5027.
90. 90 Castro-Castro, A., et al. ATAT1/MEC-17 acetyltransferase and HDAC6 deacetylase control a balance of acetylation of alpha-tubulin and cortactin and regulate MT1-MMP trafficking and breast tumor cell invasion. Eur. J. Cell Biol. 91 (2012), 950–960.
91. 91 Kashiwaya, K., et al. Involvement of the tubulin tyrosine ligase-like family member 4 polyglutamylase in PELP1 polyglutamylation and chromatin remodeling in pancreatic cancer cells. Cancer Res. 70 (2010), 4024–4033.
92. 92 Froidevaux-Klipfel, L., et al. Septin cooperation with tubulin polyglutamylation contributes to cancer cell adaptation to taxanes. Oncotarget 6 (2015), 36063–36080.
93. 93 Kuroda, H., et al. Differential expression of glu-tubulin in relation to mammary gland disease. Virchows Arch. 457 (2010), 477–482.
94. 94 Soucek, K., et al. Normal and prostate cancer cells display distinct molecular profiles of alpha-tubulin posttranslational modifications. Prostate 66 (2006), 954–965.
95. 95 Wasylyk, C., et al. Tubulin tyrosine ligase like 12 links to prostate cancer through tubulin posttranslational modification and chromosome ploidy. Int. J. Cancer 127 (2010), 2542–2553.
96. 96 Kato, C., et al. Low expression of human tubulin tyrosine ligase and suppressed tubulin tyrosination/detyrosination cycle are associated with impaired neuronal differentiation in neuroblastomas with poor prognosis. Int. J. Cancer 112 (2004), 365–375.
97. 97 Whipple, R.A., et al. Epithelial-to-mesenchymal transition promotes tubulin detyrosination and microtentacles that enhance endothelial engagement. Cancer Res. 70 (2010), 8127–8137.
98. 98 Roll-Mecak, A., Vale, R.D., Structural basis of microtubule severing by the hereditary spastic paraplegia protein spastin. Nature 451 (2008), 363–367.