The roles of tubulin post-translational modifications in cancer

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The roles of tubulin post-translational modifications in cancer

INSTITUTO DE TECNOLOGIA QUÍMICA E BIOLÓGICA ANTÓNIO XAVIER| UNIVERSIDADE NOVA DE LISBOA

Danilo da Silva Lopes

Dissertation presented to obtain the Ph.D degree in

Oeiras, February, 2023

Molecular Biosciences

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Danilo da Silva Lopes

Dissertation presented to obtain the Ph.D degree in Molecular Biosciences

Instituto de Tecnologia Química e Biológica António Xavier | Universidade NOVA de Lisboa

Oeiras, February, 2023

post-translational

modifications in cancer

Research work coordinated by:

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Orientador: Doutor Helder José Martins Maiato

Chromosome Instability and Dynamics, i3s - Instituto de Investigação e Inovacão em Saúde, Universidade do Porto, Rua Alfredo Allen 208, 4200-135 Porto, Portugal.

Departamento de Biomedicina, Faculdade de Medicina, Universidade do Porto, Alameda Prof.

Hernâni Monteiro, 4200-319 Porto, Portugal.

Coorientadora: Doutora Mónica Bettencourt-Dias

Cell Cycle Regulation, IGC - Instituto Gulbenkian de Ciência, Rua da Quinta Grande 6, 2780-156 Oeiras, Portugal.

PhD supported by a fellowship from Fundação para a Ciência e a Tecnologia of Portugal (SFRH/BD/135077/2017) and the Graduate Program Science for Development (PGCD).

Oeiras, February, 2023

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ACKNOWLEDGMENTS

The completion of this thesis was only possible with the help of many people.

Therefore, here I express my sincere acknowledgments.

To my supervisor Helder Maiato, for the opportunity and privilege of working in his lab and understand the beauty of the dividing cell. Since I started, Helder challenged me, gave me the freedom to explore, and showed me how exciting science can be. Many thanks for always showing me that my limits were beyond what I thought, and that resilience is extremely important in science. Your daily advices, support and inspiration played an important role in the fulfillment of this goal.

To Bernardo Orr, for the supervising during the initial years. Your guidance and knowledge shared were essential to boost my work. My deepest gratitude!

To all CID lab members and alumni, for creating the best possible atmosphere for scientific work. My special thanks to Alex, Ana Almeida, Ana Figueiredo, Ariana, Carolina, Danica, Elias, Filipe, Hugo, Joana L., Joana M., Joana O., Jorge, Liam, Luisa, Marco, Margarida D., Margarida G., Naoyuki, Sonia, Toze and Vanessa. All of you made different, but extremely important contributions (scientific and nonscientific) throughout my days in the lab. You allowed me to have an easier path to fulfilling my goals. Each of you is a source of inspiration.

To Mónica Dias for accepting the co-supervision, insightful discussions and advice over these years.

To my thesis committee, Elsa Logarinho and Joana Paredes for the essential guidance and advice during this Ph.D.

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To all i3s community, that directly or indirectly contributed to this journey.

Special thanks to the help of the i3s facilities, where many experiments were performed.

To PGCD, for the opportunity of this Ph.D. It was such a pleasure to be part of this innovative and inspiring Ph.D. program. Special thanks to Joana Sá, Patricia Beldade, Carla Semedo, Inês Maciel and Leonor Ruivo for the advices and support. To my PGCD colleagues, for the friendship and company during the first year of Ph.D. I will always carry this community with me! To ITQB NOVA, especially the academic office for the bureaucratic help.

To Fundação para a Ciência e a Tecnologia of Portugal for the support through a fellowship.

To all my friends who supported me and were always fundamental in my life.

To Hercules, who always had to listen to my boring explanations about cell biology, and Kaori, for the coffee therapy chats. A special thanks to Porto´s gang, Deisy, Miguel, Irina, Baltazar and JC, for their friendship, scientific help, as well as the great moments, parties and dinners, which made this journey easier.

À minha família, especialmente meus pais e meus irmãos pelo apoio e por ser uma fonte de força para aguentar a distância. Minhas irmãs Kelisa e Dulce, pela constante preocupação e palavras de encorajamento. Ao meu irmão Kevin, o primeiro a me mostrar o encanto que é a biologia; obrigado por teres sido uma fonte de inspiração.

À minha mãe, a minha inspiração, meu exemplo de vida e de resiliência.

Muito obrigado pelo amor incondicional. Cheguei até aqui porque sempre me apoiaste, incentivaste a correr atrás de quaisquer que fossem os meus sonhos e acreditaste em mim.

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Por fim, à Julie, por ser o meu principal apoio e meu amor. Obrigado por fortemente me apoiares e entrar comigo nesta aventura intercontinental.

Também, por estares ao meu lado em cada momento deste doutoramento, tanto nos difíceis e em cada vitória. É imensurável a minha gratidão pela tua paciência e pelo teu incentivo diário a tudo o que faço, como se fosse teu.

Gracias por compartir tu vida conmigo y por tanto!

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ABSTRACT

Microtubules are dynamic structures of the cytoskeleton composed by several α/β-tubulin isotypes that may undergo various post-translational modifications (PTMs). This “tubulin code” generates microtubule diversity that modulates key cellular functions. However, whether and how these tubulin PTMs regulate cancer cell properties remains largely unknown. Here, we proposed to understand: (1) the regulation of tubulin PTMs in cancer; (2) the functional implications of the “cancer tubulin code”. For these purposes, we performed a comprehensive analysis and dissection of tubulin PTMs in the NCI-60 cancer cell panel. Strikingly, α-tubulin acetylation, detyrosination and related ∆2 modification, which typically accumulate on stable microtubules, varied significantly and were found frequently uncoupled among different cancer cells. Interestingly, high α-tubulin acetylation associated with, but was not required for, cancer cell cytotoxicity to taxol/paclitaxel. On the other hand, experimental increase of α-tubulin detyrosination enhanced taxol cytotoxicity by promoting cell death in mitosis and in the subsequent interphase. Mechanistically, depletion of MCAK, a microtubule-depolymerizing enzyme inhibited by α-tubulin detyrosination, led to similar outcomes, which were not cumulative with increased α-tubulin detyrosination. Curiously, only increased α-tubulin detyrosination aggravated taxol-induced spindle multipolarity. Thus, α-tubulin detyrosination promotes taxol cytotoxicity by suppressing MCAK activity, likely by promoting chromosome missegregation, regardless of spindle defects. Lastly, we investigated the implication of α-tubulin detyrosination for metastatic behavior. Experimental increase of this PTM generated changes in cell morphology and cell-cell interactions that promoted epithelial-to- mesenchymal transition, suggestive of the initiation of metastasis. Overall, this thesis provided an initial dissection of the cancer tubulin code, and

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identified microtubule acetylation as a predictive determinant of taxol response, while uncovering roles for microtubule detyrosination in taxol- induced cytotoxicity and metastasis.

Keywords: tubulin code, acetylation, detyrosination, MCAK, paclitaxel, chromosomal instability, tubulin, taxol, cancer

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RESUMO

Os microtúbulos são estruturas dinâmicas do citoesqueleto compostas por vários isotipos de α/β-tubulina, que podem sofrer várias modificações pós- traducionais (MPTs). Este “tubulin code” gera diversidade de microtúbulos que modulam funções celulares essenciais. No entanto, se e como esses MPTs de tubulina regulam as propriedades das células cancerígenas permanece em grande parte desconhecido. Aqui, propusemos compreender: (1) a regulação dos MPTs de tubulina no cancro; (2) as implicações funcionais do “cancer tubulin code”. Portanto, realizamos uma análise abrangente e dissecção de MPTs de tubulina no painel de células cancerígenas NCI-60. Surpreendentemente, acetilação, destirosinação e a modificação ∆2 da α-tubulina, que normalmente acumulam em microtúbulos estáveis, variaram significativamente e foram encontradas frequentemente desacopladas entre diferentes células cancerígenas. Curiosamente, alta acetilação de α-tubulina associou com, mas não foi necessária para, a citotoxicidade das células cancerígenas ao taxol/paclitaxel. Por outro lado, o aumento experimental da destirosinação de α-tubulina aumentou a citotoxicidade ao taxol, através da promoção da morte celular na mitose e na interfase subsequente. Mecanisticamente, a depleção da proteína MCAK, uma enzima despolimerizante de microtúbulos inibida pela destirosinação da α-tubulina, levou a resultados semelhantes, que não foram cumulativos com o aumento da destirosinação da α-tubulina.

Curiosamente, apenas o aumento da destirosinação da α-tubulina agravou a multipolaridade do fuso mitótico induzida pelo taxol. Portanto, a destirosinação de α-tubulina promove a citotoxicidade ao taxol através da supressão da atividade de MCAK, provavelmente promovendo a incorreta segregação cromossómica, independentemente de defeitos do fuso mitótico. Por fim, investigamos a implicação da destirosinação da α-tubulina

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no comportamento metastático. O aumento experimental deste MPT gerou alterações na morfologia celular e nas interações célula-célula que promoveram a transição epitélio-mesenquimal, sugestiva do início da metástase. No geral, esta tese forneceu uma dissecção inicial do “cancer tubulin code” e identificou a acetilação dos microtúbulos como um determinante preditivo da resposta ao taxol, ao mesmo tempo em que revela papéis da destirosinação de microtúbulos na citotoxicidade induzida por taxol e metástase.

Palavras-chave: tubulin code, acetilação, destirosinação, MCAK, paclitaxel, instabilidade cromossómica, tubulina, taxol, cancro

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LIST OF FIGURES

CHAPTER 1

Figure 1.1. The phases and regulators of the eukaryotic cell cycle. ... 4

Figure 1.2. The stages of mitosis in eukaryotic cells. ... 5

Figure 1.3. Microtubule organization and dynamics. ... 9

Figure 1.4. The tubulin code. ... 12

Figure 1.5. Summary of the established roles of the tubulin code in mitosis. ... 16

Figure 1.6. Modes of kinetochore-microtubule attachments. ... 17

Figure 1.7. Implications of the tubulin code for tumor progression and metastasis. ... 26

Figure 1.8. Paclitaxel (Taxol®) binding site and mechanisms of action. ... 31

CHAPTER 2 Figure 2.1.1. Immunoblot screen of tubulin PTMs in the NCI-60 cancer cell panel. ... 53

Figure 2.1.2. Cancer cells display highly variable tubulin PTM signatures. 55 Figure 2.1.3. Tubulin PTMs only vary slightly throughout the cell cycle. .... 55

Figure 2.1.4. α-tubulin detyrosination, Δ2 and acetylation can be uncoupled in cancer cells. ... 57

Figure 2.1.5. VASH2 expression weakly correlates with α-tubulin detyrosination, while VASH1 and MATCAP weakly correlates with TTL levels. ... 60

Figure 2.1.6. High α-tubulin acetylation correlates with taxol cytotoxicity. . 62

Figure 2.1.7. α-tubulin detyrosination, Δ2, TTL and MCAK levels do not correlate with taxol cytotoxicity, whereas α-tubulin detyrosination and TTL do not correlate with MCAK. ... 63

Figure 2.1.8. α-tubulin acetylation levels upon different experimental manipulations and characterization of α-tubulin PTMs in human U2OS cells. ... 65

Figure 2.1.9. α-tubulin acetylation does not interfere with taxol cytotoxicity. ... 66

Figure 2.1.11. α-tubulin acetylation does not affect microtubule dynamics. ... 70

Figure 2.1.12. Microtubule detyrosination promotes taxol cytotoxicity mainly by regulating MCAK activity. ... 73

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II Figure 2.1.13. Increased α-tubulin detyrosination or MCAK depletion

enhances taxol cytotoxicity in T-47D cells, but does not revert taxol-

resistance in OVCAR-4 cells. ... 74 Figure 2.1.14. Increased α-tubulin detyrosination, but not MCAK depletion, aggravates taxol-induced mitotic spindle multipolarity. ... 78 Figure 2.1.15. High α-tubulin detyrosination or MCAK depletion promotes taxol-induced cell death in mitosis and in the subsequent interphase. ... 81 Figure 2.2.1 Low α-tubulin detyrosination and high α-tubulin acetylation correlates with increased cell crowding. ... 95 Figure 2.2.2. Excessive α-tubulin detyrosination induces cell disassociation and mesenchymal phenotype. ... 97 Figure 2.2.3. α-tubulin detyrosination increases the size of active focal adhesion at the cell edge. ... 98

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III

LIST OF TABLES

Table 1. Tubulin isotypes, post-translational modifications and modifying enzymes in cancer ... 22 Table 2. Summary of the NCI-60 cancer cell lines used in the study. ... 40

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IV

SCIENTIFIC PUBLICATIONS

Lopes, D., Seabra, A. L., Orr, B., Maiato, H. α-tubulin detyrosination links the suppression of MCAK activity with taxol cytotoxicity. Journal of Cell Biology, 2022. DOI: 10.1083/jcb.202205092

Lopes, D., & Maiato, H. The tubulin code in mitosis and cancer. Cells, 2020.

DOI: 10.3390/cells9112356.

Ferreira, L. T., Figueiredo, A. C., Orr, B., Lopes, D., Maiato, H. Dissecting the role of the tubulin code in mitosis. Methods in Cell Biology, 2018. DOI:

10.1016/bs.mcb.2018.03.040.

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1

CHAPTER 1. GENERAL INTRODUCTION

(This chapter contains parts of the review: Lopes, D., & Maiato, H. The tubulin code in mitosis and cancer. Cells, 2020. DOI: 10.3390/cells9112356.)

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2 1.1 The cell cycle

Since the first description of cells, observed in cork by Robert Hooke in 1665 (Hooke, 1665), the understanding of cell biology attracted the interest of scientists over centuries. Indeed, in early 1850s, Robert Remak proposed that cells are derived from other cells (Remak, 1855). This theory of cell multiplication was made popular by Rudolf Virchow (Virchow, 1858), thus opening the field of cell division. In Eukaryotes, a sequence of events occurs during the cell cycle to duplicate the genetic material of the cell, followed by its division into two identical daughter cells. S phase (in interphase) and M phase (mitosis) are the two major phases that respectively promotes DNA replication and chromosome segregation during cell division.

1.1.1 Cell cycle stages

Eukaryotic cells follow four sequential phases of a cycle that allow them to growth, synthesize their DNA and other components, and divide. Known as the longer period of the cell cycle, interphase includes G1, S and G2 phases.

After mitosis, the Gap 1 phase (G1) is the longer phase, which allows cell growth and prepares for DNA replication. Depending on extracellular signals, cells in G1 can exit the cycle and enter G0, a resting state in which they do not proliferate. This state can be reverted and cells re-enter the cell cycle in G1 phase. During S phase (synthesis of DNA), the genome is replicated into two identical copies. After this process is completed, cells proceed to G2, a second Gap phase where they continue to grow and prepare for DNA segregation. Finally, an essential shorter phase, known as mitosis, takes place, when cells equally distribute the duplicated DNA content and divide into two daughters (reviewed in (Alberts et al., 2017; Vermeulen et al., 2003)) (Figure 1.1).

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3 The progression of the cell cycle depends on the regulation and activity of specific proteins that allow the transition between phases. Cyclin- dependent kinases (Cdks) are a family of protein kinases that phosphorylate substrates essential for cell cycle control. The activity of these enzymes is regulated by the interaction with cyclins, activating proteins whose expression oscillates throughout the cell cycle. Thus, during cell cycle progression, Cdk levels remain normal while their activities vary between phases (Martinez-Alonso and Malumbres, 2020; Vermeulen et al., 2003).

Cyclin D activates Cdk4 and Cdk6, promoting cell cycle entry and G1 progression (Martinez-Alonso and Malumbres, 2020; Sherr, 1993). In order to promote DNA replication, Cyclin D is degraded and the formation of Cyclin E-Cdk2 complex is required for G1/S transition (Ohtsubo et al., 1995). Cyclin A expression is essential in two phases of cell cycle. The complex Cyclin A- Cdk2 control DNA synthesis during S phase, while Cyclin A-Cdk1 mediates G2 progression and preparation for mitosis (Pagano et al., 1992). During mitosis, the activation of the complex Cyclin B-Cdk1 is fundamental in regulating chromosome segregation (Gavet and Pines, 2010) (Figure 1.1).

(figure legends in the next page)

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4 Figure 1.1. The phases and regulators of the eukaryotic cell cycle.

Interphase is the longest phase of the cell cycle, divided into three sequential stages (G1, S, and G2), where cellular components are synthesized and DNA duplicated. At M phase, the replicated genetic content is segregated between daughter cells. The cyclin-Cdk complexes involved in different biological processes of the cycle are indicated in each stage.

1.2 Mitosis

The main purpose of cell division is to divide the replicated genetic material into two daughter cells, during a process known as mitosis. The terminology

“mitosis” (Greek: “mitos” = thread + modern Latin “osis” = process) was coined and first described in the late 1800s by Walther Flemming. Analyzing Salamander cells, Flemming successfully observed a few cytoplasmic components and nuclear stainable structures (“chromatin”) by using haematein and haematoxylin dyes. The mother cell chromatin rearranged into threadlike structures (chromosomes), which moved to the center of the cell and then separated generating two new daughter nuclei (Flemming, 1882). Later, taking advantage of polarized light and electron microscopy, chromosome movement was associated to the fibrous structures of the mitotic spindle, currently known as microtubules (Brinkley and Stubblefield, 1966; Inoue, 1953).

Over more than a century, as interest in understanding mitosis increased, contributions of many scientists allowed a detailed organization into five sequential phases: prophase, prometaphase, metaphase, anaphase and telophase (Figure 1.2). The beginning of mitosis, known as prophase, is marked by the condensation of the duplicated chromosomes and centrosome separation to opposite poles, where microtubules emanate, starting the building of a bipolar spindle (Pines and Rieder, 2001). Usually

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5 not emphasized as a phase of mitosis, prometaphase starts upon nuclear envelope breakdown (NEB), leading to interaction between spindle microtubules and chromosomes at their kinetochores, thus promoting chromosome movement to the spindle equator (Ferreira and Maiato, 2021).

The cell is only considered in metaphase when all chromosomes align at the equator (Ferreira and Maiato, 2021; Maiato et al., 2017). During anaphase, sister chromatids segregate to opposite poles through pulling forces exerted by microtubule depolymerization and sliding of anti-parallel microtubules (McIntosh, 2021; Vukusic and Tolic, 2021). Throughout telophase, chromosomes at the poles begin to decondense along with nuclear envelope reformation (NER), generating two daughter nuclei (Pines and Rieder, 2001).

Concomitant with the last stages of mitosis, during cytokinesis, the cytoplasm of the parental cell is divided into two daughter cells, which also separates the two nuclei (Eggert et al., 2006) (Figure 1.2).

Figure 1.2. The stages of mitosis in eukaryotic cells.

Prophase – chromosomes start to condensate and the centrosomes move to opposite poles. Prometaphase – after nuclear envelope breakdown (NEB), microtubules growing from the poles capture the chromosomes promoting its congression to the cell equator. Metaphase - the chromosomes reach the alignment at the spindle equator. Anaphase – chromosomes segregate through a poleward movement. Telophase – marks the end of mitosis, with sister chromatid decondensation and nuclear envelope reformation (NER) of the two daughter nuclei. Cytokinesis gives rise to two daughter cells.

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6 1.3 Microtubules

Microtubules (MTs) are main structures of the cytoskeleton found in all eukaryotic cells. They are dynamic, hollow cylinders, typically formed by protofilaments of α/β-tubulin heterodimers that are involved in key cellular processes, such as cell division, motility, differentiation and intracellular transport.

During the late 1930s and early 1950s, polarized light microscopy allowed the first identification of microtubules (Inoue, 1953; Schmidt, 1939).

A decade later, with electron microscopes, these “fibrils” of the mitotic spindle were better analyzed (Roth and Daniels, 1962),leading to the detailed identification and description of these filaments as hollow tubular structures, which were named as “microtubules” (Ledbetter and Porter, 1963;

Slautterback, 1963). Few years later, tubulin, the main structural component of microtubules, was identified by its capacity to associate with colchicine(Borisy and Taylor, 1967a; Borisy and Taylor, 1967b)., with the term “tubulin” introduced soon after (Mohri, 1968).

1.3.1 Structure

Microtubules are generally formed by 13 protofilaments, composed by α/β- tubulin (55 kDa each) heterodimers of 8 nm (4 nm each unit) (Amos and Klug, 1974), that laterally associate forming a tubular structure of 25 nm diameter (Desai and Mitchison, 1997; Ledbetter and Porter, 1964; Nogales et al., 1998). α/β-tubulin heterodimers align in a longitudinal head-to-tail association, which generates the polarity of microtubules. While α-tubulin is oriented toward the minus end, β-tubulin is at the plus end, with a faster polymerizing activity (Fan et al., 1996; Mitchison, 1993; Nogales et al., 1998) (Figure 1.3). Additionally, γ-tubulin, another member of the tubulin family (Oakley and Oakley, 1989), is localized at microtubule organizing centers

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7 (MTOCs) (Horio et al., 1991). As part of γ-TuRC (γ-tubulin ring complex), γ- tubulin is essential for microtubule nucleation, through interaction with α- and β-tubulin (Kollman et al., 2011).

α-tubulin and β-tubulin are structurally similar, containing three functional domains: the nucleotide-binding N-terminal, an intermediate, and a C-terminal domain (Nogales et al., 1998). Therefore, these monomers bind to guanosine triphosphate (GTP) with high affinity. Although GTP is not exchangeable at the α-tubulin binding site (N-site), it is hydrolyzed to guanosine diphosphate (GDP) at β-tubulin binding site (E-site) (Mandelkow and Mandelkow, 1989; Mitchison, 1993; Nogales et al., 1998).

1.3.2 Microtubule dynamics

The understanding of microtubules as dynamic structures started in the early 1970s with the first in vitro polymerization studies of microtubules (Weisenberg, 1972). In 1984 Mitchison and Kirschner observed that microtubules have cycles of growth and shrinkage, which they termed dynamic instability (Mitchison and Kirschner, 1984a; Mitchison and Kirschner, 1984b). Following studies confirmed this process in vivo (Cassimeris et al., 1988; Sammak and Borisy, 1988; Sammak et al., 1987).

Microtubule polymerization and depolymerization cycles are based on GTP hydrolysis. The soluble GTP-bound tubulin heterodimers assemble, and the hydrolysis of the β-tubulin GTP occurs in the microtubule lattice, generating GDP-tubulin heterodimers (Desai and Mitchison, 1997). In addition, the following incorporated GTP dimers form a cap that stabilizes the microtubule (GTP cap model). Under the decreasing cap, the unstable GDP lattice is exposed, starting a rapid depolymerization (catastrophe) that ends with the microtubule disassembly or the addition of new GTP subunits (rescue), thus recapping the polymer (Gudimchuk and McIntosh, 2021;

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8 Mitchison and Kirschner, 1984a) (Figure 1.3). During shrinkage, protofilaments lose their linearity and start bending, which destabilizes the microtubule. Interestingly, this conformational change may be explained by two models: the allosteric and lattice models. The first proposes that GTP hydrolysis generates curved GDP-tubulin dimers, thereby increasing the tension between neighbors in the microtubule lattice, leading to microtubule depolymerization. On the other hand, the second model suggests that the strength of the lateral and longitudinal bonds, or the flexibility of the GDP- tubulin dimers are affected (Gudimchuk and McIntosh, 2021).

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9 Figure 1.3. Microtubule organization and dynamics.

(A) Microtubules are composed by α/β-tubulin heterodimers of 8nm diameter (4 nm each unit), that organize longitudinally into protofilaments. While α- tubulin points toward the minus end, β-tubulin faces the plus end. (B) A microtubule is a hollow cylinder of 25 nm diameter, formed by lateral interaction of thirteen protofilaments. (C) In a growing microtubule, GTP- tubulin heterodimers are incorporated to the microtubule plus end, followed by their hydrolysis in the lattice. These recently added dimers form a GTP cap that stabilizes the microtubule. Additionally, a microtubule in a paused state of growing can be reached, which may undergo additional growth or start depolymerization. However, after GTP hydrolysis, the protofilaments start bending and the microtubule shrinks. Adapted from (Akhmanova and Steinmetz, 2008).

1.4 The tubulin code

α- and β-tubulin proteins are encoded by several different genes (also known as tubulin isotypes) that diverge in their C-terminal tail regarding length and amino acid composition (Figure 1.4) (Janke, 2014; Ludueña and Banerjee, 2008). In eukaryotes, the expression and distribution of different tubulin isotypes is cell- and tissue-specific (Ludueña and Banerjee, 2008). In addition, α- and β-tubulin isotypes may undergo multiple post-translational modifications (PTMs). As α/β-tubulin heterodimers polymerize into microtubules, the combination of isotype expression with PTMs generates microtubule diversity or a “tubulin code” (Figure 1.4), which has been implicated in the regulation of microtubule properties and functions underlying fundamental cellular processes (Janke and Magiera, 2020;

Verhey and Gaertig, 2007).

Acetylation, detyrosination, polyglutamylation and polyglycylation are amongst the best characterized tubulin PTMs (Figure 1.4). Acetylation occurs in both α- and β-tubulins, more specifically at the luminal-side Lysine-

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10 40 (K40) of α-tubulin (L'Hernault and Rosenbaum, 1985; Soppina et al., 2012) and Lysine 252 (K252) of β-tubulin (Chu et al., 2011). While K252 is modified by the acetyltransferase San (Chu et al., 2011), K40 is acetylated by the acetyltransferase MEC-17/αTAT1 (Akella et al., 2010; Shida et al., 2010) and deacetylated by histone deacetylase 6 (HDAC6) and sirtuin2 (SIRT2) (Hubbert et al., 2002; North et al., 2003). When incorporated into microtubules, α-tubulin can also be detyrosinated, which consists on the catalytic removal of the last tyrosine present at the C-terminal tail of most isoforms by tubulin carboxypeptidases (TCPs), including the recently identified MATCAP (Landskron et al., 2022), and Vasohibin 1 (VASH1) and Vasohibin 2 (VASH2) complexes with their associated Small Vasohibin- Binding Protein (SVBP) (Aillaud et al., 2017; Li et al., 2019; Li et al., 2020;

Liao et al., 2019; Liu et al., 2019; Nieuwenhuis et al., 2017; Wang et al., 2019). As microtubules depolymerize, soluble detyrosinated α-tubulin can be retyrosinated by a highly specific tubulin tyrosine ligase (TTL) that closes the cycle (Ersfeld et al., 1993; Schroder et al., 1985). After detyrosination, α- tubulin C-terminal tails may also be subject to the removal of the penultimate and antepenultimate glutamates by cytosolic carboxypeptidases (CCPs) (Rogowski et al., 2010; Tort et al., 2014), leading to formation of the non- tyrosinatable Δ2- and Δ3-tubulin, respectively (Aillaud et al., 2016; Paturle- Lafanechere et al., 1991). Additionally, C-terminal tails of both α- and β- tubulins undergo side-chain polyglutamylation and polyglycylation (Edde et al., 1990; Redeker et al., 1994). The single or consecutive addition of glutamate residues to the -carboxyl group of C-terminal tails is performed by several TTL-like (TTLL) (poly)glutamylases (Janke and Magiera, 2020;

Janke et al., 2005; van Dijk et al., 2007) and is/are removed by a set of CCPs known as deglutamylases (Janke and Magiera, 2020; Kimura et al., 2010;

Rogowski et al., 2010; Tort et al., 2014). Similarly, the addition of glycine residues relies on the (poly)glycylases TTLL3, TTLL8 and TTLL10 (Rogowski

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11 et al., 2009; Wloga et al., 2009), but the identity of tubulin deglycylases remains unknown. Lastly, several other tubulin PTMs, such as methylation, polyamination, phosphorylation, ubiquitinylation, sumoylation, palmitoylation (reviewed in (Janke and Magiera, 2020)) and O-GlcNAcylation (Walgren et al., 2003) occur in the tubulin core structure adjacent to the C-terminal tails.

These PTMs remain poorly characterized at the functional level but are likely to be implicated in microtubule assembly and dynamics (Janke and Magiera, 2020; Ji et al., 2011; Tian and Qin, 2019).

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12 Figure 1.4. The tubulin code.

The tubulin code combines different tubulin isotypes and post-translational modifications (PTMs) to generate microtubule diversity. Only the best- characterized isotypes and PTMs (+ respective enzymes) are depicted. See main text for details. Adapted from (Lopes and Maiato, 2020)

1.5 The tubulin code in mitosis

Mitosis relies on the critical contribution of microtubules, as well as several microtubule-associated proteins (MAPs) and motors, to regulate several key mechanisms underlying the faithful segregation of the genetic material during cell division. It involves the assembly of a specialized microtubule-based structure known as the mitotic spindle. Due to their intrinsic dynamic nature, mitotic spindle microtubules are vastly tyrosinated, i.e., remain essentially nonmodified (note that most gene-encoded α-tubulin isoforms carry a last Tyrosine residue at their C-terminal tails; see Figure 1.4). As some spindle microtubules become gradually stabilized due to the establishment of chromosome attachments at the kinetochore, as well as possible interactions between some interpolar microtubules, they become increasingly detyrosinated (Barisic et al., 2015; Ferreira et al., 2020; Gundersen and Bulinski, 1986; Gundersen et al., 1984; Liao et al., 2019; Peris et al., 2006) (Figure 1.5). Likewise, kinetochore microtubules are highly acetylated on the K40 of α-tubulin (Barisic et al., 2015; Wilson and Forer, 1989b), polyglutamylated (Bobinnec et al., 1998b) and accumulated Δ2-tubulin (Ferreira et al., 2018). The actions of spindle microtubules during mitosis is regulated by several MAPs (Maiato et al., 2004) and assisted by several motor proteins (Cross and McAinsh, 2014). For instance, the initial capture and transport of peripheral chromosomes by microtubules is mediated by dynein/dynactin (Hayden et al., 1990; Li et al., 2007; Vorozhko et al., 2008;

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13 Yang et al., 2007), a minus-end-directed motor localized at unattached kinetochores (Pfarr et al., 1990; Steuer et al., 1990), whereas the subsequent congression to the spindle equator is mediated by another kinetochore- associated motor, Centromere Protein E (CENP-E)/kinesin-7, with microtubule plus-end-directed activity (Kapoor et al., 2006; Wood et al., 1997). Other mitotic motors include kinesin-5, which slides antiparallel microtubules to ensure proper centrosome separation, spindle bipolarity and spindle elongation during anaphase, as well as kinesin-13s, which lack motor activity but promote microtubule depolymerization to control spindle length and mediate mitotic error correction (Andrews et al., 2004; Bakhoum et al., 2009b; Domnitz et al., 2012; Kline-Smith et al., 2004; Lan et al., 2004; Mann and Wadsworth, 2019). Thus, the mitotic spindle is an anisotropic and highly heterogeneous structure, with dynamic astral microtubules essentially tyrosinated, in contrast with more stable microtubule subpopulations, such as kinetochore and a fraction of interpolar microtubules, which accumulate detyrosinated, Δ2, acetylated and polyglutamylated tubulin. How these modifications impact the action of the different mitotic motors that assist chromosome segregation remains poorly understood.

1.5.1 A navigation system guides chromosomes to the spindle equator Although tubulin diversity in the mitotic spindle has been recognized for several decades, the respective functional relevance for mitosis remained unclear until recently. One crucial implication of the tubulin code hypothesis is the regulation of MAPs and motors by specific tubulin isotypes and PTMs (Verhey and Gaertig, 2007). Original work in neurons revealed that classic kinesin motors, such as Kinesin-1, are able to recognize and have a preference for microtubules with particular tubulin PTMs, namely detyrosination and acetylation (Konishi and Setou, 2009; Reed et al., 2006).

Subsequently, -tubulin detyrosination was shown to regulate mitotic

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14 chromosome congression to the metaphase plate by guiding the microtubule plus-end-directed motor CENP-E/kinesin-7 at kinetochores in human cells (Barisic et al., 2015). In contrast, the microtubule minus-end-directed motor dynein/dynactin that is also localized at unattached kinetochores (Pfarr et al., 1990; Steuer et al., 1990), preferentially associates with tyrosinated microtubules (McKenney et al., 2014; McKenney et al., 2016; Nirschl et al., 2016; Peris et al., 2006), which favor the initiation of motion, but are dispensable for subsequent dynein/dynactin processivity (McKenney et al., 2016; Nirschl et al., 2016). Thus, detyrosinated/tyrosinated -tubulin regulates the activity of opposing kinetochore motors, establishing a navigation system for chromosomes that assists their congression to the spindle equator (Barisic and Maiato, 2016) (Figure 1.5). Accordingly, during the initial capture of chromosomes, dynein/dynactin counteracts the action of chromokinesins on chromosome arms to move peripheral chromosomes along tyrosinated astral microtubules towards the vicinity of the poles (Barisic et al., 2014). By transporting peripheral chromosomes to the poles where the microtubule destabilizing activity of Aurora A kinase is higher (Chmatal et al., 2015; Ye et al., 2015), dynein/dynactin prevents the formation of stable end- on kinetochore–microtubule attachments that would otherwise cause the random ejection of polar chromosomes by chromokinesins (Barisic et al., 2014; Barisic and Maiato, 2016). Once at the poles, Aurora A-mediated phosphorylation activates CENP-E at kinetochores of polar chromosomes (Kim et al., 2010), thus allowing their transport specifically along detyrosinated spindle microtubules towards the equator. In agreement, recent super-resolution coherent-hybrid stimulated emission depletion microscopy (Pereira et al., 2019) of CENP-E-GFP revealed its exclusive association with stable kinetochore and interpolar microtubule bundles but not with tyrosinated astral microtubules (Steblyanko, 2020). Curiously, - tubulin acetylation on K40, which is also enriched on stable spindle

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15 microtubules (Wilson and Forer, 1989b), does not interfere with polar chromosome congression (Barisic et al., 2015). While the potential contribution of other tubulin PTMs to chromosome congression remains unknown, these findings support a robust working model in which tyrosinated/detyrosinated microtubules guide peripheral chromosomes towards the spindle equator.

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16 Figure 1.5. Summary of the established roles of the tubulin code in mitosis.

The initial capture of peripheral chromosomes is mediated by dynein/dynactin at kinetochores, upon which, the chromosome is brought to the vicinity of the centrosome by lateral transport along tyrosinated astral microtubules. This prevents the random ejection of the chromosome by the action of Chromokinesins on chromosome arms. Once at the pole, high Aurora A activity prevents the stabilization of end-on kinetochore-microtubule attachments, which otherwise would favor the action of Chromokinesins on chromosome arms. In parallel, Aurora A-mediated phosphorylation activates CENP-E at kinetochores. This initiates transport towards the spindle equator (congression) along stable detyrosinated microtubules. Mitotic centromere- associated kinesin (MCAK) and Kif2b (not depicted) at centromeres and kinetochores are also inhibited by tubulin detyrosination on kinetochore microtubules, allowing the correction of syntelic and merotelic attachments, while preserving correct amphitelic attachments on bi-oriented chromosomes. MCAK at the microtubule plus ends also regulates astral microtubule length to allow interaction with dynein/dynactin at the cortex or cytoplasmic organelles (not depicted), which exerts pulling forces necessary for spindle orientation and positioning. See main text for details. (Lopes and Maiato, 2020)

1.5.2 The tubulin code in error correction 1.5.2.1 Error correction

The regulation of kinetochore microtubule dynamics is essential for error correction and the maintenance of genome stability, since it allows the establishment of correct attachments of the spindle microtubules to the chromosomes. Known as amphitelic (correct) attachments, sister kinetochores are attached to microtubules from opposite spindle poles, leading to chromosome bi-orientation. Under erroneous kinetochore- microtubule attachments, both sister kinetochores can be oriented towards a single spindle pole (syntelic), and/or a single kinetochore can be attached

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17 with microtubules oriented to both poles (merotelic) and/or only a sister kinetochore attached to a single pole (monotelic) (reviewed in (Tanaka, 2010)) (Figure 1.6).

Figure 1.6. Modes of kinetochore-microtubule attachments.

Monotelic – a single sister kinetochore attached to one pole. Syntelic – both kinetochores attached to the same pole. Amphitelic – sister kinetochores attached to opposite poles. Merotelic – a single sister kinetochore attached to both poles. Adapted from (Tanaka, 2010).

1.5.2.2 Kinesin-13s

Kinesin superfamily 2a (Kif2a), kinesin superfamily 2b (Kif2b) and mitotic centromere-associated kinesin (MCAK) are members of the kinesin-13 family and microtubule depolymerizers, involved in several cellular functions such as spindle assembly, chromosome positioning and error correction (Walczak et al., 2013). The best characterized member, MCAK is a powerful microtubule depolymerizer, composed by four domains: N-terminal, a positively charged neck domain, the catalytic domain containing the microtubule and ATP binding sites, and lastly a C-terminal domain (reviewed

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18 in (Ritter et al., 2015; Wordeman, 2005)). To depolymerize microtubules, MCAK in solution is in a “closed” conformation and binds to microtubules through the microtubule binding site. Then, ATP is hydrolyzed and MCAK changes to the “open” conformation, which allows its sliding through the microtubule until it reaches the end. At the end, the interaction between the tubulin dimer and MCAK´s KVD motif leads to a conformational change into a catalytic competent MCAK, which starts depolymerization. Lastly, a second ATP hydrolysis occurs that releases the tubulin dimer bound to MCAK, generating an “open” conformation MCAK prepared for the next cycle (reviewed in (Ritter et al., 2015)).

Kinesin-13s, such as Kif2b and MCAK, promote kinetochore microtubule dynamics, thus playing a key role in the correction of mal- oriented chromosomes with erroneous kinetochore-microtubule attachments and, ultimately, in the prevention of chromosome missegregation (Bakhoum et al., 2009a; Bakhoum et al., 2009b) (Figure 1.5). In agreement, stimulation of kinetochore microtubule dynamics in otherwise chromosomally unstable cancer cells by increasing kinesin-13 depolymerase activity reestablished chromosomal stability (Bakhoum et al., 2018; Bakhoum et al., 2009b).

1.5.2.3 A mitotic error code

Building on the previous finding that MCAK´s microtubule depolymerizing activity is reduced four-fold in the presence of detyrosinated microtubules in vitro (Peris et al., 2009; Sirajuddin et al., 2014), it was recently shown that the mitotic error correction activity of MCAK and Kif2b is regulated by α- tubulin detyrosination (Ferreira et al., 2020). Accordingly, the experimental depletion of TTL or overexpression of VASH1-SVBP, which caused a constitutive increase of α-tubulin detyrosination in the vicinity of the kinetochores, compromised error correction, leading to chromosome segregation errors. Importantly, α-tubulin detyrosination specifically impaired

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19 the MCAK-based error correction machinery located on centromeres/kinetochores, and it did so without affecting global kinetochore microtubule dynamics, suggesting that mitotic error correction is exquisitely sensitive to the detyrosinated state of α-tubulin that likely occurs at the individual microtubule level. These data support the existence of a “mitotic error code” in which α-tubulin detyrosination/tyrosination signals and regulates MCAK activity at centromeres/kinetochores to discriminate between correct and incorrect kinetochore-MT attachments during mitosis (Figure 1.5).

Complete centrosome separation before nuclear envelope breakdown prevents subsequent segregation errors and ensures mitotic fidelity (Silkworth et al., 2012). This relies on several elements, including the microtubule motors kinesin-5, required for centrosome separation, and dynein/dynactin, which promote both centrosome separation and positioning (Nunes et al., 2020; Raaijmakers et al., 2012). Similar to dynein/dynactin, kinesin-5 appears to have increased affinity to tyrosinated dendritic microtubules in neurons (Kahn et al., 2015), but direct evidence from in vitro reconstitution assays is still lacking. Nonetheless, recent work in which centrosome positioning in human mitotic cells was tracked in 3D indicated that centrosome separation at nuclear envelope breakdown is insensitive to the tyrosinated state of -tubulin (Ferreira et al., 2020). This reinforces the idea that the observed increase in mitotic errors associated with excessive α-tubulin detyrosination is due to the incapacity to correct, rather than an increased propensity to make errors.

1.5.3 Role in mitotic spindle orientation and positioning

Mitotic spindle orientation and positioning in the cell center is essential for accurate cell division and relies on the action of pulling forces on astral microtubules (Siller and Doe, 2009). In particular, dynein/dynactin anchored

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20 to cortical proteins or cytoplasmic organelles was shown to play a significant role in spindle orientation/positioning (Barbosa et al., 2017; Kotak et al., 2012; Nguyen-Ngoc et al., 2007), possibly through its increased affinity to tyrosinated astral microtubules (Figure 1.5). Indeed, modulation of α-tubulin tyrosination state, either through TTL knockout (Peris et al., 2006) or CRISPR/Cas9-mediated editing of the C-terminal tyrosine (Barbosa et al., 2017), caused spindle orientation defects. In contrast, an experimental decrease of α-tubulin detyrosination after VASH1/2 silencing increased the depolymerase activity of MCAK, resulting in disoriented spindles, with shorter astral microtubules (Liao et al., 2019). Taken together, these observations indicate that the mechanisms behind spindle orientation/positioning rely on the intrinsic nature (i.e., nonmodified) of tyrosinated α-tubulin to allow astral microtubules to establish a correct cell division plane (Figure 1.5).

1.5.4 Roles in centrosome structure and cytokinesis

Tubulin polyglutamylation is highly enriched on centriole microtubules (Bobinnec et al., 1998a; Bobinnec et al., 1998b) and has been proposed to contribute to normal mitosis by maintaining centrosome structure (Abal et al., 2005; Bobinnec et al., 1998a). Indeed, recent super-resolution imaging of centriole structure revealed the specific distribution of polyglutamylation on centriole MTs and suggested a key role for this PTM in ultrastructural organization of specific centriolar proteins (Mahecic et al., 2020).

Furthermore, tubulin polyglutamylation promotes the activity of the microtubule-severing enzymes spastin and katanin (Lacroix et al., 2010;

Sharma et al., 2007; Shin et al., 2019; Valenstein and Roll-Mecak, 2016), which are also implicated in cell division. Indeed, their activities regulate several cellular processes that likely impact chromosome segregation fidelity, such as microtubule poleward flux, spindle orientation and length (Jiang et al., 2017; McNally et al., 2006; Zhang et al., 2007a). Spastin and

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21 katanin are also required for the abscission step and completion of cytokinesis (Connell et al., 2009; Guizetti et al., 2011; Matsuo et al., 2013).

Like spastin (Connell et al., 2009) and katanin (Matsuo et al., 2013), polyglutamylated tubulin is enriched at the midbody (Lacroix et al., 2010), and a tubulin mutation that compromises polyglutamylation (and, possibly, also polyglycylation) in cilia was shown to cause cytokinesis defects (Thazhath et al., 2002). These results suggest that the completion of cytokinesis relies on the regulation of spastin and katanin activities by tubulin polyglutamylation.

1.6 The cancer tubulin code

1.6.1 (De)Regulation of tubulin isotypes and PTMs in cancer

Several works have reported an emerging link between alterations of tubulin isotypes and PTMs and/or associated modifying enzymes with certain cancers; most noticeable, those occurring in breast, colon, prostate, liver, brain, bile duct and pancreas (Table 1). These alterations often correlate with specific cancer properties, including poor outcome/prognosis (Boggs et al., 2015; Kato et al., 2004; Mialhe et al., 2001) and metastatic ability (Boggs et al., 2015), supporting the potential use of cancer tubulin isotypes and/or PTM signatures as useful biomarkers, as well as for therapeutic purposes.

However, a comprehensive and definitive view on the real potential is still lacking, especially concerning causality, since the available data is still limited and often contradictory.

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22 Table 1. Tubulin isotypes, post-translational modifications and modifying enzymes in cancer

Tubulin PTM (and/or

Enzymes)/ Isotype Cancer Regulation References

Detyrosination

Prostate Cancer Cells Upregulated (Soucek et al., 2006) Poor Prognosis Breast

Tumors Upregulated (Mialhe et al., 2001) Invasive Ductal

Carcinoma (Breast) Upregulated (Whipple et al., 2010)

TTL

Prostate Cancer Cells Downregulated (Soucek et al., 2006) Poor Prognosis

Neuroblastomas Downregulated (Kato et al., 2004) VASH2

Hepatocellular carcinoma Tissues and

Cell Lines

Upregulated (Xue et al., 2013) Δ2-Tubulin Prostate Cancer Cells Downregulated (Soucek et

al., 2006) Acetylation Metastatic Breast

Tumors and Cell Lines Upregulated (Boggs et al., 2015)

HDAC6

Pancreatic Tumors Upregulated (Li et al., 2014) Glioblastoma Tissues

and Cell Lines Upregulated (Wang et al., 2016) Cholangiocarcinoma

Cell Lines Upregulated (Gradilone et al., 2013) Glutamylation/

Polyglutamylation Prostate Cancer Cells Upregulated (Soucek et al., 2006) TTLL4 Pancreatic Ductal

Adenocarcinoma Cells Upregulated (Kashiwaya et al., 2010) Glycylation

TTLL3

Colon Tumors and Cell

Lines Downregulated (Rocha et al., 2014)

β3-tubulin

Pancreatic Tumors and

Cell Lines Upregulated (McCarroll et al., 2015) Pancreatic Ductal

Adenocarcinoma Tissues

Upregulated (Lee et al., 2007) Breast Cancer Brain

Metastases Upregulated (Kanojia et al., 2015)

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23 1.6.2 Functional implications of the cancer tubulin code

The differential regulation of specific tubulin isotypes and/or PTMs in cancer might reflect their role in key mechanisms underlying cell transformation (Figure 1.7). β3-tubulin (TUBB3) is the most frequent tubulin isotype associated with specific cancer features. Its expression was proposed to be important for tumor development (McCarroll et al., 2010; McCarroll et al., 2015) and metastatic ability (Kanojia et al., 2015; McCarroll et al., 2015), correlating with poor outcomes (Ferrandina et al., 2006; Kanojia et al., 2015) and response to microtubule-targeting agents commonly used in chemotherapy (reviewed in (Parker et al., 2017)). The expression of other isotypes such as β2-tubulin, and its altered cellular localization in colorectal cancer, also correlate with poor outcomes (Ruksha et al., 2019).

In addition, the regulation of cell proliferation, which is essential for cancer development, was proposed to be mediated by certain tubulin PTMs.

In this regard, the tubulin glycylase TTLL3 was proposed to restrict cell proliferation in the colon and is downregulated in colon cancer (Rocha et al., 2014), whereas the tubulin glutamylase TTLL4 was suggested to promote cell proliferation in pancreatic cancer cells (Kashiwaya et al., 2010).

However, whether this was specifically due to a role of TTLL4 in tubulin glutamylation remains controversial, since an additional activity towards non- tubulin substrates has been reported (Regnard et al., 2000; van Dijk et al., 2008). The tubulin acetyltransferase αTat1 was also shown to be required for contact inhibition of cell proliferation in vitro (Aguilar et al., 2014). In agreement, the tubulin deacetylase HDAC6 seems to promote cell proliferation in several cancer cell lines (Gradilone et al., 2013; Lee et al., 2008; Putcha et al., 2015; Wang et al., 2016; Woan et al., 2015), consistent with its upregulation in some cancers (Table 1). Nevertheless, specificity remains to be demonstrated, since HDAC6 is also known to modulate

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24 acetylation of other substrates besides tubulin (Zhang et al., 2007b).

Interestingly, the activity of TTLL3 and HDAC6 was also proposed to impact tumorigenesis. Accordingly, the experimental loss of TTLL3 in a mouse model of tumorigenesis resulted in the development of cancer, thus validating its downregulation in colon cancer and suggesting a cancer- suppressing role for tubulin glycylation (Rocha et al., 2014). In contrast, the expression of HDAC6 promoted colony and spheroid formation of cancer cells, as well as tumor growth in mice (Gradilone et al., 2013; Lee et al., 2008;

Wang et al., 2016; Woan et al., 2015). The activity of other tubulin-modifying enzymes, such as TTL, is also decreased during tumorigenesis in mouse models, resulting in increased detyrosinated- and Δ2-tubulin levels (Lafanechere et al., 1998). This is consistent with the association between

-tubulin detyrosination and tumor aggressiveness (Mialhe et al., 2001), as well as with the frequent downregulation of TTL and consequent upregulation of -tubulin detyrosination in several cancers (Table 1).

The recent discovery of Vasohibins (VASH1 and VASH2) as TCPs (Aillaud et al., 2017; Nieuwenhuis et al., 2017) revitalized the discussion about the role of tubulin detyrosination in cancer. Vasohibins and their associated SVBP were originally identified as secreted proteins implicated in angiogenesis (Sato, 2013). While VASH2 promotes vascularity by accumulating at the sprouting zone, VASH1 expression is increased in endothelial cells of the termination zone, where it inhibits vascularity (Kimura et al., 2009). During tumor development in mice xenograft models, experiments involving administration of ectopic VASH1 indicated that it inhibits tumor lymphangiogenesis (Heishi et al., 2010), angiogenesis and growth (Hosaka et al., 2009). On the other hand, VASH2, which appears to play an important role in cancer cell proliferation (Xue et al., 2013), promotes tumor angiogenesis and growth (Kitahara et al., 2014; Koyanagi et al., 2013;

Takahashi et al., 2012; Xue et al., 2013). Noteworthy, none of these studies

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25 demonstrate that the observed impact in cancer was due to defective tubulin detyrosination. However, human patients suffering from a broad range of carcinomas had mutations in VASH1 and VASH2 that compromised their tubulin detyrosination activity (Wang et al., 2019) and, more recently, it was suggested that the MT detyrosinating activity of VASH1 inhibited angiogenesis by interfering with endocytosis and trafficking of proangiogenic factor receptors (Kobayashi et al., 2021). Taken together, these findings suggest that, in addition to the downregulation of TTL (Lafanechere et al., 1998), the link between tubulin detyrosination and tumorigenesis may be attributed to the role of Vasohibins in angiogenesis. The availability of VASH1/2-SVBP knockout mice (Kimura et al., 2009; Pagnamenta et al., 2019) will be instrumental in clarifying the apparently opposite roles of VASH1 and VASH2 in cancer and whether this is due to their secreted and/or tubulin detyrosinating activities.

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26 Figure 1.7. Implications of the tubulin code for tumor progression and metastasis.

While the downregulation of TTLL3 (glycylation), together with the expression of VASH2 (detyrosination), HDAC6 (acetylation) and β3-tubulin, promotes tumor growth, this is inhibited by VASH1 (detyrosination). Tumor formation and chromosomal instability is also associated with the downregulation of TTL. Tubulin acetylation, detyrosination and β3-tubulin isotypes might promote several steps of metastasis associated with the epithelial-to-mesenchymal transition, such as cell migration and invasion.

See main text for details. (Lopes and Maiato, 2020)

1.6.3 The cancer tubulin code in cell migration and invasion

Tubulin PTMs have also been implicated in epithelial-to-mesenchymal transition (EMT), a key process behind metastasis initiation. For instance, experimental increase of the tubulin deacetylase HDAC6 promoted EMT, whereas TGF-β induction of EMT downregulated tubulin acetylation (Gu et al., 2016). Likewise, the induction of EMT also correlated with the downregulation of TTL and the consequent increase of tubulin detyrosination (Whipple et al., 2010), as shown before during tumor development (Lafanechere et al., 1998), thus pointing to the possible involvement of these tubulin PTMs and associated enzymes in cell transformation.

Interestingly, tubulin acetylation is also frequently associated with the regulation of cell migration, although this remains controversial. While HDAC6 expression and activity was proposed to promote cell migration (Bance et al., 2019; Gu et al., 2016; Haggarty et al., 2003; Hubbert et al., 2002; Zhang et al., 2007b), the opposite effect was observed after the loss of αTat1 or mutation of the α-tubulin lysine 40 (K40R) (Bance et al., 2019;

Boggs et al., 2015; Castro-Castro et al., 2012; Montagnac et al., 2013). The establishment of cell adhesion to the substrate also has implications for cell motility, and the loss of either HDAC6 or αTat1 leads to an increased focal

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27 adhesion area and number, respectively, as well as decreased dynamics (Bance et al., 2019; Tran et al., 2007). However, other works reported that loss of αTat1 leads to a decrease in focal adhesion number (Aguilar et al., 2014). The basis for this discrepancy remains unclear, but it is likely associated with different experimental setups; one study investigated the role of αTat1 in wound-induced migrating cells (Bance et al., 2019), while the other used normally growing cells (Aguilar et al., 2014), raising the possibility that αTat1 promotes focal adhesion dynamics specifically during cell migration.

The upregulation of tubulin acetylation in metastatic breast tumors and cell lines (Boggs et al., 2015) is consistent with its association with cancer cell invasiveness. RNAi-mediated depletion of either αTat1 or HDAC6 indicated that their expression induced breast cancer cell invasion (Castro- Castro et al., 2012; Montagnac et al., 2013; Rey et al., 2011). Additionally, the increased tubulin acetylation of these metastatic breast cancer cells promoted micro-tentacle generation and cell reattachment ability, essential for metastasis (Boggs et al., 2015). Likewise, a high frequency of micro- tentacles and cell reattachment were also associated with tubulin detyrosination (Whipple et al., 2010; Whipple et al., 2013). Collectively, these data favor a potential role of tubulin acetylation in metastasis progression.

While HDAC6 indiscriminately acts upon multiple protein targets, the direct modulation of tubulin acetylation by K40R mutation experiments suggest that the upregulation of tubulin acetylation is a metastasis-promoting factor, supporting the αTat1-related findings. This would explain the link between the upregulation of tubulin acetylation and poor prognosis in breast cancer patients (Boggs et al., 2015), but unspecific effects due to the overexpression of GFP-tagged K40R mutant α-tubulin cannot be excluded.

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28 1.6.4 How alterations of the tubulin code in mitosis might be implicated

in cancer

Chromosomal instability, a hallmark of cancers, has been shown to promote the metastatic process (Bakhoum et al., 2018). Indeed, the overexpression of Kif2b or MCAK, in addition to reestablishing the stability of chromosomally unstable cancer cells (Bakhoum et al., 2018; Bakhoum et al., 2009b), inhibits metastasis in vitro and in vivo, with a consequent increase in survival (Bakhoum et al., 2018). Given that excessive tubulin detyrosination might lead to chromosomal instability by suppressing the error correction activity of MCAK and Kif2b (Ferreira et al., 2020), together with the observed upregulation of tubulin detyrosination in invasive cancer and with poor prognosis (Table 1), it raises the exciting possibility that an increase in tubulin detyrosination might promote cancer progression through inhibition of the mitotic error correction machinery. However, an extensive analysis of tubulin detyrosination in chromosomal instability-prone cancers, together with the elucidation of its implications for cancer metastasis, is necessary for its establishment as potential diagnostic and prognostic biomarkers. In addition, tubulin detyrosination represents a promising therapeutic target for cancer suppression-for example, by using TCP inhibitors, such as epoY (Aillaud et al., 2017) or parthenolide (Fonrose et al., 2007). This might implicate in the activity of microtubule-targeting agents, such as taxol, which resistance in cells is affected by modulation of MCAK (Ganguly et al., 2011a; Ganguly et al., 2011b).

The deregulation of tubulin detyrosination in cancers might also be involved in other mitotic-related cancer features. Firstly, the cell cycle delay observed upon VASH1/2 (Liao et al., 2019) and VASH2 (Xue et al., 2013) deletion might unveil the importance of VASH2 for proper cancer cell proliferation and tumor development (Kitahara et al., 2014; Koyanagi et al.,

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29 2013; Takahashi et al., 2012; Xue et al., 2013). Furthermore, both experimental upregulation and downregulation of tubulin detyrosination led to congression defects, causing alterations in CENP-E-mediated transport of chromosomes to the spindle equator (Barisic et al., 2015). Additionally, the decrease of CENP-E expression is well-established to promote mild chromosomal instability and aneuploidy, as well as tumorigenesis in mice (Clemente-Ruiz et al., 2014; Silk et al., 2013; Weaver et al., 2007; Zasadil et al., 2016). Therefore, the deregulation of tubulin detyrosination in cancers (Table 1) may also account for cancer promotion under conditions of moderate chromosomal instability, such as those associated with mild problems in chromosome congression. Further investigation is required to fully understand the potential implications of tubulin detyrosination and other PTMs for tumorigenesis and the respective link with chromosomal instability.

1.7 Microtubule-targeting agents

Microtubules are extremely important in several cellular processes, making them one of the main targets for diverse chemotherapeutic agents.

Microtubule-targeting agents have broad natural origins, from a diversity of plants and animals, and upon binding to microtubules alter their polymerization and dynamics, which ultimately may lead to cell death (reviewed in (Dumontet and Jordan, 2010; Jordan and Wilson, 2004;

Kavallaris, 2010)). According to their effect, these compounds can be separated into two groups: microtubule-stabilizing or microtubule- destabilizing agents. The stabilizing agents promote microtubule assembly by binding to the taxane (paclitaxel, docetaxel, epothilones) or laulimalide/peloruside (laulimalide, peloruside A) binding site on β-tubulin.

On the other hand, destabilizing agents bind to the vinca (vinca alkaloids, eribulin), colchicine (colchicine, nocodazole, combretastatins), maytansine, or pironetin site on α- or β-tubulin, which induces microtubule

The roles of tubulin post-translational modifications in cancer