細胞微管在調控各種細胞活動中扮演了重要的角色，過去雖然能使用抗微管藥物針對微管進行研究，但由於抗微管藥物的作用速度極慢，且無法針對特定的微管進行精準的破壞，因此無法有效地用於研究微管與細胞的動態調控機制，為了突破這樣的技術限制，我們改造了微管切割蛋白Spastin，使其能受化學遺傳學技術調控，精準的聚集到特定的微管群上進行切割，同時整個反應又具備可逆性，利用這項技術，我們能分別專一的破壞初級纖毛的軸絲、紡錘絲，以及胞間橋。透過這項技術，我們觀察到急劇的破壞微管會迅速的終止膜囊泡與溶酶體的運輸、引發高基氏體與內質網型態重組；並在不影響粒線體膜電位的情況下，減緩粒線體的融合與分裂；破壞微管也會促進細胞形成收縮的應力纖維，進而導致細胞的硬度上升。這項技術提供了我們一個新的角度來研究細胞微管是如何調控細胞構造及功能。

Microtubules tightly regulate various cellular activities. Our understanding of microtubules is largely based on microtubule-targeting agents, which, however, are insufficient to dissect the dynamic mechanisms of specific microtubule populations due to their slow effects on the entire pool of microtubules. To address this limitation, we have incorporated chemogenetics to disassemble specific microtubule subtypes including ciliary axonemes, mitotic spindles, and intercellular bridges, by rapidly recruiting engineered microtubule-cleaving enzymes onto target microtubules. Acute microtubule disassembly swiftly halted vesicular trafficking and lysosomal dynamics. It also immediately triggered Golgi and ER reorganization and slowed the fusion/fission of mitochondria without affecting mitochondrial membrane potential. Cell rigidity was increased after microtubule disruption owing to increased contractile stress fibers. These tools enable to uncover new insights of how microtubules precisely regulate cellular architectures and functions.
This thesis is modified from my previously published work in EMBO Journal, titled 'Precise control of microtubule disassembly in living cells'.

Chapter 1 Introduction 1
1.1 Overview of microtubules 1
1.2 Endogenous processes to regulate MT dynamics 1
1.3 Overview of the microtubule severing enzyme, Spastin 2
1.4 Overview of present microtubule disruption techniques 3
Chapter 2 Methods and Materials 6
2.1 Cell culture and transfection 6
2.2 DNA construction 6
2.3 Immunofluorescence staining 7
2.4 Western blotting 8
2.5 Live-cell imaging 9
2.6 Measurement of microtubule filament area 10
2.7 Tracking vesicles and lysosomes 10
2.8 Analysis of mitochondrial morphology and dynamics 11
2.9 Cell synchronization 12
2.10 Statistical analysis 12
2.11 List of antibodies and dyes 13
2.12 List of chemical reagents 14
2.13 Supplementary sequences 15
Chapter 3 Results 20
3.1 Rapid translocation of proteins of interest onto microtubules 20
3.2 Engineering microtubule-severing enzymes for precise microtubule disruption 21
3.3 Rapid disruption of specific microtubule-based structures 24
3.4 Acute microtubule disassembly inhibits vesicular trafficking and lysosomal dynamics 26
3.5 Microtubules are essential for organization of Golgi and endoplasmic reticulum 27
3.6 Microtubules are essential for the dynamics of mitochondria 28
3.7 Acute microtubule disassembly immediately triggers stress fiber formation and increases cell rigidity 31
Chapter 4 Discussion and Conclusion 34
References 38
Figures 50
Figure 1. Rapid translocation of proteins of interest onto microtubules 50
Figure 2. The microtubule-severing activity of engineered Spastin enzymes 53
Figure 3. Rapidly recruiting engineered Spastins onto microtubules 55
Figure 4. The microtubule disruption rates of MTAs and our system 57
Figure 5. Our system can disrupt long-lived acetylated microtubules 59
Figure 6. Rapid disassembly of microtubules in COS7 cells 61
Figure 7. Rapid disassembly of microtubules in U2OS cells 63
Figure 8. Using a gibberellin-based system to rapidly translocate proteins of interest onto microtubules 65
Figure 9. Using the gibberellin-based system to rapidly disassemble microtubules 67
Figure 10. Acute microtubule disruption is not dependent on proteasome-mediated degradation 69
Figure 11. Inhibition of Spastin activity reverses microtubule disassembly 71
Figure 12. Disassembly rates of detyrosinated and hyperglutamylated microtubules 73
Figure 13. Subcellular distribution of CFP-FRB-MAP4m at different cell cycle phases 76
Figure 14. Rapid disassembly of ciliary axonemes 78
Figure 15. The structure of the primary cilia occasionally does not collapse after axoneme disassembly 80
Figure 16. Rapid disassembly of mitotic spindles and intercellular bridges 82
Figure 17. Spastin does not rapidly disrupt centrosomes 84
Figure 18. Acute microtubule disassembly halts vesicular trafficking 86
Figure 19. Acute microtubule disassembly halts lysosomal dynamics 88
Figure 20. Acute microtubule disassembly reorganizes the Golgi apparatus 90
Figure 21. Acute microtubule disassembly reorganizes the ER 91
Figure 22. The effects of acute microtubule disassembly on the dynamics and activities of mitochondria 92
Figure 23. Microtubule disassembly enhances stress fiber formation and cell rigidity 95
Figure 24. Method used to measure microtubule filament area in real time 98

1. Liu, G. Y. et al. Precise control of microtubule disassembly in living cells. EMBO J. 41, (2022).
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).
3. Cleary, J. M. & Hancock, W. O. Molecular mechanisms underlying microtubule growth dynamics. Curr. Biol. 31, R560–R573 (2021).
4. Yang, W. T. et al. The Emerging Roles of Axonemal Glutamylation in Regulation of Cilia Architecture and Functions. Frontiers in Cell and Developmental Biology vol. 9 (2021).
5. Prosser, S. L. & Pelletier, L. Mitotic spindle assembly in animal cells: A fine balancing act. Nat. Rev. Mol. Cell Biol. 18, 187–201 (2017).
6. Antanavičiūtė, I., Gibieža, P., Prekeris, R. & Skeberdis, V. A. Midbody: From the regulator of cytokinesis to postmitotic signaling organelle. Med. 54, 1–10 (2018).
7. Doxsey, S., McCollum, D. & Theurkauf, W. Centrosomes in cellular regulation. Annu. Rev. Cell Dev. Biol. 21, 411–434 (2005).
8. Hildebrandt, F., Benzing, T. & Katsanis, N. Ciliopathies. N. Engl. J. Med. 364, 1533–1543 (2011).
9. Sferra, A., Nicita, F. & Bertini, E. Microtubule dysfunction: A common feature of neurodegenerative diseases. Int. J. Mol. Sci. 21, 1–36 (2020).
10. Parker, A. L., Kavallaris, M. & McCarroll, J. A. Microtubules and their role in cellular stress in cancer. Front. Oncol. 4 JUN, 1–19 (2014).
11. Akhmanova, A. & Steinmetz, M. O. Control of microtubule organization and dynamics: Two ends in the limelight. Nat. Rev. Mol. Cell Biol. 16, 711–726 (2015).
12. Roll-Mecak, A. & McNally, F. J. Microtubule-severing enzymes. Curr. Opin. Cell Biol. 22, 96–103 (2010).
13. Allison, R., Edgar, J. R. & Reid, E. Spastin MIT Domain Disease-Associated Mutations Disrupt Lysosomal Function. Front. Neurosci. 13, 1–14 (2019).
14. Sharp, D. J. & Ross, J. L. Microtubule-severing enzymes at the cutting edge. J. Cell Sci. 125, 2561–2569 (2012).
15. Roll-Mecak, A. & Vale, R. D. Making more microtubules by severing: A common theme of noncentrosomal microtubule arrays? J. Cell Biol. 175, 849–851 (2006).
16. Roll-Mecak, A. & Vale, R. D. The Drosophila homologue of the hereditary spastic paraplegia protein, spastin, severs and disassembles microtubules. Curr. Biol. 15, 650–655 (2005).
17. Ghosh, D. K., Dasgupta, D. & Guha, A. Models, Regulations, and Functions of Microtubule Severing by Katanin. ISRN Mol. Biol. 2012, 1–14 (2012).
18. Hazan, J. et al. Spastin, a new AAA protein, is altered in the most frequent form of autosomal dominant spastic paraplegia. Nat. Genet. 23, 296–303 (1999).
19. Claudiani, P., Riano, E., Errico, A., Andolfi, G. & Rugarli, E. I. Spastin subcellular localization is regulated through usage of different translation start sites and active export from the nucleus. Exp. Cell Res. 309, 358–369 (2005).
20. Solowska, J. M. et al. Quantitative and functional analyses of spastin in the nervous system: Implications for hereditary spastic paraplegia. J. Neurosci. 28, 2147–2157 (2008).
21. Solowska, J. M., Rao, A. N. & Baas, P. W. Truncating mutations of SPAST associated with hereditary spastic paraplegia indicate greater accumulation and toxicity of the M1 isoform of spastin. Mol. Biol. Cell 28, 1728–1737 (2017).
22. White, S. R., Evans, K. J., Lary, J., Cole, J. L. & Lauring, B. Recognition of C-terminal amino acids in tubulin by pore loops in Spastin is important for microtubule severing. J. Cell Biol. 176, 995–1005 (2007).
23. Han, H. et al. Structure of spastin bound to a glutamate-rich peptide implies a hand-over-hand mechanism of substrate translocation. J. Biol. Chem. 295, 435–443 (2020).
24. Jessop, M., Felix, J. & Gutsche, I. AAA+ ATPases: structural insertions under the magnifying glass. Curr. Opin. Struct. Biol. 66, 119–128 (2021).
25. Eckert, T., Le, D. T. Van, Link, S., Friedmann, L. & Woehlke, G. Spastin’s Microtubule-Binding Properties and Comparison to Katanin. PLoS One 7, 1–16 (2012).
26. Roll-Mecak, A. & Vale, R. D. Structural basis of microtubule severing by the hereditary spastic paraplegia protein spastin. Nature (2008) doi:10.1038/nature06482.
27. Steinmetz, M. O. & Prota, A. E. Microtubule-Targeting Agents: Strategies To Hijack the Cytoskeleton. Trends Cell Biol. 28, 776–792 (2018).
28. Rovini, A., Savry, A., Braguer, D. & Carré, M. Microtubule-targeted agents: When mitochondria become essential to chemotherapy. Biochim. Biophys. Acta - Bioenerg. 1807, 679–688 (2011).
29. LeDizet, M. & Piperno, G. Cytoplasmic microtubules containing acetylated α-tubulin in Chlamydomonas reinhardtii: Spatial arrangement and properties. J. Cell Biol. 103, 13–22 (1986).
30. Xu, Z. et al. Microtubules acquire resistance from mechanical breakage through intralumenal acetylation. Science (80-. ). 356, 328–332 (2017).
31. Xie, R., Nguyen, S., McKeehan, W. L. & Liu, L. Acetylated microtubules are required for fusion of autophagosomes with lysosomes. BMC Cell Biol. 11, 89 (2010).
32. Friedman, J. R., Webster, B. M., Mastronarde, D. N., Verhey, K. J. & Voeltz, G. K. ER sliding dynamics and ER-mitochondrial contacts occur on acetylated microtubules. J. Cell Biol. 190, 363–375 (2010).
33. Molero, J. C., Whitehead, J. P., Meerloo, T. & James, D. E. Nocodazole Inhibits Insulin-stimulated Glucose Transport in 3T3-L1 Adipocytes via a Microtubule-independent Mechanism. J. Biol. Chem. 276, 43829–43835 (2001).
34. Huby, R. D. J., Weiss, A. & Ley, S. C. Nocodazole inhibits signal transduction by the T cell antigen receptor. J. Biol. Chem. 273, 12024–12031 (1998).
35. Bates, D. & Eastman, A. Microtubule destabilising agents: far more than just antimitotic anticancer drugs. Br. J. Clin. Pharmacol. 83, 255–268 (2017).
36. Van Haren, J. et al. Local control of intracellular microtubule dynamics by EB1 photodissociation. Nat. Cell Biol. 20, 252–261 (2018).
37. Adikes, R. C., Hallett, R. A., Saway, B. F., Kuhlman, B. & Slep, K. C. Control of microtubule dynamics using an optogenetic microtubule plus end-F-actin cross-linker. J. Cell Biol. 217, 779–793 (2018).
38. Borowiak, M. et al. Photoswitchable Inhibitors of Microtubule Dynamics Optically Control Mitosis and Cell Death. Cell 162, 403–411 (2015).
39. Valente, A. J., Maddalena, L. A., Robb, E. L., Moradi, F. & Stuart, J. A. A simple ImageJ macro tool for analyzing mitochondrial network morphology in mammalian cell culture. Acta Histochem. 119, 315–326 (2017).
40. Lefebvre, A. E. Y. T., Ma, D., Kessenbrock, K., Lawson, D. A. & Digman, M. A. Automated segmentation and tracking of mitochondria in live-cell time-lapse images. Nat. Methods (2021) doi:10.1038/s41592-021-01234-z.
41. Derose, R., Miyamoto, T. & Inoue, T. Manipulating signaling at will: Chemically-inducible dimerization (CID) techniques resolve problems in cell biology. Pflugers Arch. Eur. J. Physiol. 465, 409–417 (2013).
42. Fan, C.-H. H. et al. Manipulating cellular activities using an ultrasound-chemical hybrid tool. ACS Synth. Biol. 6, acssynbio.7b00162 (2017).
43. Hong, S. R. et al. Spatiotemporal manipulation of ciliary glutamylation reveals its roles in intraciliary trafficking and Hedgehog signaling. Nat. Commun. 9, 1–13 (2018).
44. Rodrigues, T. B. & Ballesteros, P. Spastin and Microtubules: Functions in Health and Disease. J. Neurosci. Res. 3253, 3244–3253 (2007).
45. Lacroix, B. et al. Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. J. Cell Biol. 189, 945–954 (2010).
46. Cupido, T., Pisa, R., Kelley, M. E. & Kapoor, T. M. Designing a chemical inhibitor for the AAA protein spastin using active site mutations. Nat. Chem. Biol. 15, 444–452 (2019).
47. Aillaud, C. et al. Vasohibins/SVBP are tubulin carboxypeptidases (TCPs) that regulate neuron differentiation. Science (80-. ). 1453, 1448–1453 (2017).
48. van Dijk, J. et al. A Targeted Multienzyme Mechanism for Selective Microtubule Polyglutamylation. Mol. Cell 26, 437–448 (2007).
49. Gillingham, A. K. & Munro, S. The PACT domain, a conserved centrosomal targeting motif in the coiled-coil proteins AKAP450 and pericentrin. EMBO Rep. 1, 524–529 (2000).
50. Schuh, M. An actin-dependent mechanism for long-range vesicle transport. Nat. Cell Biol. 13, 1431–1436 (2011).
51. Ba, Q., Raghavan, G., Kiselyov, K. & Yang, G. Whole-Cell Scale Dynamic Organization of Lysosomes Revealed by Spatial Statistical Analysis. Cell Rep. 23, 3591–3606 (2018).
52. Noordstra, I. & Akhmanova, A. Linking cortical microtubule attachment and exocytosis. F1000Research 6, 1–12 (2017).
53. 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).
54. Jaqaman, K. et al. Robust single-particle tracking in live-cell time-lapse sequences. Nat. Methods 5, 695–702 (2008).
55. Guo, Y. et al. Visualizing Intracellular Organelle and Cytoskeletal Interactions at Nanoscale Resolution on Millisecond Timescales. Cell 175, 1430-1442.e17 (2018).
56. Terasaki, M., Chen, L. B. & Fujiwara, K. Microtubules and the endoplasmic reticulum are highly interdependent structures. J. Cell Biol. 103, 1557–1568 (1986).
57. Bannai, H., Inoue, T., Nakayama, T., Hattori, M. & Mikoshiba, K. Kinesin dependent, rapid, bi-directional transport of ER sub-compartment in dendrites of hippocampal neurons. J. Cell Sci. 117, 163–175 (2004).
58. Shibata, Y., Hu, J., Kozlov, M. M. & Rapoport, T. A. Mechanisms shaping the membranes of cellular organelles. Annu. Rev. Cell Dev. Biol. 25, 329–354 (2009).
59. Poteryaev, D., Squirrell, J. M., Campbell, J. M., White, J. G. & Spang, A. Involvement of the Actin Cytoskeleton and Homotypic Membrane Fusion in ER Dynamics in Caenorhabditis elegans. Mol. Biol. Cell 16, 2139–2153 (2005).
60. Lu, L., Ladinsky, M. & Kirchhausen, T. Cisternal Organization of the Endoplasmic Reticulum during Mitosis. Mol. Biol. Cell 20, 3471–3480 (2009).
61. Schroeder, L. K. et al. Dynamic nanoscale morphology of the ER surveyed by STED microscopy. J. Cell Biol. 218, 83–96 (2019).
62. Westermann, B. Mitochondrial fusion and fission in cell life and death. Nat. Rev. Mol. Cell Biol. 11, 872–884 (2010).
63. Schwarz, T. L. Mitochondrial trafficking in neurons. Cold Spring Harb. Perspect. Med. 3, 1–16 (2013).
64. Murota, Y., Tabu, K. & Taga, T. Requirement of ABC transporter inhibition and Hoechst 33342 dye deprivation for the assessment of side population-defined C6 glioma stem cell metabolism using fluorescent probes. BMC Cancer 16, 1–7 (2016).
65. Csordás, G. et al. Imaging Interorganelle Contacts and Local Calcium Dynamics at the ER-Mitochondrial Interface. Mol. Cell 39, 121–132 (2010).
66. Zheng, P. et al. DNA damage triggers tubular endoplasmic reticulum extension to promote apoptosis by facilitating ER-mitochondria signaling. Cell Res. 28, (2018).
67. Pimm, M. L. & Henty-Ridilla, J. L. New twists in actin-microtubule interactions. Mol. Biol. Cell 32, 211–217 (2021).
68. Dogterom, M. & Koenderink, G. H. Actin–microtubule crosstalk in cell biology. Nat. Rev. Mol. Cell Biol. 20, 38–54 (2019).
69. Kuo, C. H. et al. VEGF-Induced Endothelial Podosomes via ROCK2-Dependent Thrombomodulin Expression Initiate Sprouting Angiogenesis. Arterioscler. Thromb. Vasc. Biol. 41, 1657–1671 (2021).
70. Dou, Y., Arlock, P. & Arner, A. Blebbistatin specifically inhibits actin-myosin interaction in mouse cardiac muscle. Am. J. Physiol. - Cell Physiol. 293, 1148–1153 (2007).
71. Cojoc, D. et al. Properties of the force exerted by filopodia and lamellipodia and the involvement of cytoskeletal components. PLoS One 2, (2007).
72. Inoue, T., Heo, W. Do, Grimley, J. S., Wandless, T. J. & Meyer, T. An inducible translocation strategy to rapidly activate and inhibit small GTPase signaling pathways. Nat. Methods 2, 415–418 (2005).
73. Waterman-Storer, C. M., Worthylake, R. A., Liu, B. P., Burridget, K. & Salmon, E. D. Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat. Cell Biol. 1, 45–50 (1999).
74. Valenstein, M. L. & Roll-Mecak, A. Graded Control of Microtubule Severing by Tubulin Glutamylation. Cell 164, 911–921 (2016).
75. Lacroix, B. et al. Tubulin polyglutamylation stimulates spastin-mediated microtubule severing. J. Cell Biol. 189, 945–954 (2010).
76. Kuo, Y. W., Trottier, O., Mahamdeh, M. & Howard, J. Spastin is a dual-function enzyme that severs microtubules and promotes their regrowth to increase the number and mass of microtubules. Proc. Natl. Acad. Sci. U. S. A. 116, 5533–5541 (2019).
77. Yu, W. et al. The microtubule-severing proteins spastin and katanin participate differently in the formation of axonal branches. Mol. Biol. Cell 19, (2008).
78. Basnet, N. et al. Direct induction of microtubule branching by microtubule nucleation factor SSNA1. Nat. Cell Biol. 20, 1172–1180 (2018).
79. Errico, A., Claudiani, P., D’Addio, M. & Rugarli, E. I. Spastin interacts with the centrosomal protein NA14, and is enriched in the spindle pole, the midbody and the distal axon. Hum. Mol. Genet. 13, 2121–2132 (2004).
80. Wang, J. T., Kong, D., Hoerner, C. R., Loncarek, J. & Stearns, T. Centriole triplet microtubules are required for stable centriole formation and inheritance in human cells. bioRxiv 110, 1–17 (2017).
81. Borowiak, M. et al. Photoswitchable Inhibitors of Microtubule Dynamics Optically Control Mitosis and Cell Death. Cell 162, 403–411 (2015).
82. Müller-Deku, A. et al. Photoswitchable paclitaxel-based microtubule stabilisers allow optical control over the microtubule cytoskeleton. Nat. Commun. 11, 1–12 (2020).
83. Kesarwani, S. et al. Genetically encoded live-cell sensor for tyrosinated microtubules. J. Cell Biol. 219, (2020).