Abstract

Microtubules, composed of αβ-tubulin heterodimers, exhibit diverse structural and functional properties in different cell types. The diversity in the microtubule structure originates from tubulin heterogeneities, namely tubulin isotypes and their post-translational modifications (PTMs). These heterogeneities confer differential stability to microtubules and provide spatial cues for the functioning of the cell. Furthermore, the altered expressions of tubulin isotypes and PTMs are prominent factors for the development of resistance against some cancer drugs. In this review, we summarize our current knowledge of the tubulin isotypes and PTMs and how, together, they control the cellular functions of the microtubules. We also describe how cancer cells use this tubulin heterogeneity to acquire resistance against clinical agents and discuss existing attempts to counter the developed resistance.

Introduction

Microtubules are cylindrical polymers composed of α- and β-tubulin subunits. These subunits associate longitudinally to form protofilaments and the protofilaments associate laterally to generate a microtubule (Figure 1A) [1]. Microtubules are highly dynamic polymers as their ends can undergo rapid lengthening and shortening by the addition and removal of tubulin heterodimers, a phenomenon termed ‘dynamic instability’ [2]. Dynamic instability is characterized by two phases: ‘catastrophe’ when a microtubule shifts from a growing phase to a shortening phase, and ‘rescue’ when it returns to the state of growth from a shortening phase [3]. Microtubules also exhibit polarity as they possess two distinct ends: a more dynamic plus end, capped with β-tubulin and a less dynamic minus end with α-tubulin (Figure 1A) [3,4]. Despite their highly dynamic nature, microtubules are the most rigid filaments found in eukaryotic cells [5]. These unique properties enable the microtubules to participate in a wide variety of functions that are crucial for cellular processes such as cell division, differentiation, intracellular transport, and motility.

A model of microtubule assembly and its diverse cellular structures is shown.

Figure 1.
A model of microtubule assembly and its diverse cellular structures is shown.

(A) α-tubulin (green) and β-tubulin (blue) heterodimers associate longitudinally to form protofilaments. Thirteen protofilaments associate laterally to create a sheet, which closes to form a cylindrical microtubule. Microtubules exhibit two distinct ends; a plus end capped with β-tubulin and a minus end with α-tubulin. (B) The structurally simple microtubules can organize themselves into complicated structures to perform a wide variety of cellular functions. (i) Microtubules form the interphase microtubule array in an interphase cell, (ii) mitotic spindle in a mitotic cell, (iii) midbody in a cell undergoing cytokinesis, (iv) primary cilium in a non-motile cell, (v) motile cilia in a motile cell, (vi) flagella in a spermatozoa, (vii) marginal bands in platelets derived from a megakaryocyte and (viii) neuronal microtubule array in neurons. Microtubules in all the cells are shown in green; centrioles in red; nucleus and chromosomes in blue; and platelets in yellow with red marginal bands.

Figure 1.
A model of microtubule assembly and its diverse cellular structures is shown.

(A) α-tubulin (green) and β-tubulin (blue) heterodimers associate longitudinally to form protofilaments. Thirteen protofilaments associate laterally to create a sheet, which closes to form a cylindrical microtubule. Microtubules exhibit two distinct ends; a plus end capped with β-tubulin and a minus end with α-tubulin. (B) The structurally simple microtubules can organize themselves into complicated structures to perform a wide variety of cellular functions. (i) Microtubules form the interphase microtubule array in an interphase cell, (ii) mitotic spindle in a mitotic cell, (iii) midbody in a cell undergoing cytokinesis, (iv) primary cilium in a non-motile cell, (v) motile cilia in a motile cell, (vi) flagella in a spermatozoa, (vii) marginal bands in platelets derived from a megakaryocyte and (viii) neuronal microtubule array in neurons. Microtubules in all the cells are shown in green; centrioles in red; nucleus and chromosomes in blue; and platelets in yellow with red marginal bands.

Depending on the tissue types and functions, microtubules can form complex structures with distinct morphological and behavioral characteristics. For example, microtubules are highly dynamic in dividing cells and can undergo constant lengthening and shortening throughout the cell cycle [6]. During the interphase, microtubules nucleating from the centrosomes form an extensive network throughout the interior of the cell (Figure 1B, i). This network of microtubules facilitates intracellular transport and is crucial for the maintenance of the shape and polarity of the cell [7]. However, the relatively stable interphase microtubules undergo a complete reorganization to form a highly dynamic mitotic spindle (Figure 1B, ii) [8]. During cytokinesis, microtubules participate in the formation of midbody to facilitate the completion of cell division (Figure 1B, iii) [9]. Microtubules can also assemble into specialized structures that are involved in additional cellular functions. Axonemes are remarkably stable, nine-fold symmetric microtubule structures that form the backbone of centrioles, cilia, and flagella, which are vital for cell signaling and motility (Figure 1B, iv–vi) [10,11]. In addition to their fundamental roles in cell division, microtubules are critical for the differentiation of cells, as they can regulate the changes in cell morphology and participate in intracellular rearrangements [7,12]. The marginal bands of blood platelets consist of both stable and dynamic microtubules that maintain the discoid shape of the platelets (Figure 1B, vii) [13,14]. In neuronal cells, microtubules form a complex microtubule array required for the polarization of neurons, intracellular trafficking of cargo, neuronal migration, and differentiation (Figure 1B, viii) [15]. Although differently functioning microtubules are found in different cell types, they all are structurally similar, as they are composed of highly conserved αβ-tubulin heterodimers (Figure 1). This raises an important question of how these structurally similar microtubules exhibit diverse functionality in different cells.

Tubulin heterogeneity contributes to microtubule diversity

Earlier studies have reported the existence of subsets of microtubule populations that differ in their stabilities [6,16]. When exposed to cold, or treated with depolymerizing agents, a certain subpopulation of microtubules disassemble in minutes (t1/2 ∼ 2–5 min), whereas other microtubules exist for hours (t1/2 > 1 h), thereby being cold-resistant [16]. This difference in stability occurs as a result of the tight regulation of microtubule dynamics imparted by many microtubule-associated proteins (MAPs). MAPs, including the motor proteins, can physically associate with the microtubules and modulate the dynamic behavior of microtubules by either stabilizing or destabilizing them [17]. Apart from the regulation imparted by the MAPs, the dynamicity of the microtubules can also be regulated by direct modifications of the microtubules. These modifications can occur by two means: (1) expression of various tubulin isotypes, i.e. expression of multiple α- and β-tubulin encoding genes and (2) post-translational modifications (PTMs) of α- and β-tubulins. These modifications together confer heterogeneity to the microtubule structure, thereby contributing to its functional diversity.

Structure of microtubules

The structure of the microtubule and the sequences of α- and β-tubulin genes have been remarkably conserved in all the different cell types and organisms throughout evolution [18]. Cryo-EM analysis of the tubulin dimer revealed the presence of a structured globular domain and an unstructured negatively charged tail (Figure 2A) [19]. The N-terminal, along with the intermediate region, generates the structured core of tubulin, whereas the C-terminal forms the disordered tail [19]. During the assembly of microtubules, the globular body of tubulin stacks into protofilaments and enhances lattice formation by tubulin–tubulin interactions and the unstructured tails decorate the exterior surface of the microtubules (Figure 2B) [4]. The structured core of the tubulin has been remarkably conserved during evolution, thereby allowing very little variation in microtubule structure [18]. However, tubulin isotypes do show some sequence diversity in their core regions such as residues 55–57 and 239 in β-tubulin, which are implicated in regulating the microtubule and MAP interactions [20,21]. In contrast, tubulin isotypes exhibit significant differences in their C-terminal tails [18]. The C-terminal tail of tubulin has been proposed to be a critical spot for the regulation of microtubule dynamics due to the variations in its primary sequence and its covalent modifications (PTMs).

Arrangement of tubulin heterodimers in microtubules.

Figure 2.
Arrangement of tubulin heterodimers in microtubules.

(A) Crystal structure of the tubulin heterodimer (PDB ID: 5LYJ) is shown in cartoon representation. GTP and GDP nucleotides (orange, stick) are bound to α- and β-tubulin, respectively. Both tubulins have a magnesium atom (black, sphere) associated with them. The tubulin molecule exhibits a compactly folded globular domain and an unstructured C-terminal tail. (B) The stacking of tubulin heterodimers into a microtubule is shown in surface representation. The structured globular body of tubulin stacks to form a cylindrical microtubule, whereas the highly disordered C-terminal tails remain exposed on the exterior surface of the microtubule.

Figure 2.
Arrangement of tubulin heterodimers in microtubules.

(A) Crystal structure of the tubulin heterodimer (PDB ID: 5LYJ) is shown in cartoon representation. GTP and GDP nucleotides (orange, stick) are bound to α- and β-tubulin, respectively. Both tubulins have a magnesium atom (black, sphere) associated with them. The tubulin molecule exhibits a compactly folded globular domain and an unstructured C-terminal tail. (B) The stacking of tubulin heterodimers into a microtubule is shown in surface representation. The structured globular body of tubulin stacks to form a cylindrical microtubule, whereas the highly disordered C-terminal tails remain exposed on the exterior surface of the microtubule.

Tubulin heterogeneities

Tubulin isotypes

An earlier study performed with crane-fly and rat sperms noted that microtubules display different structures and stabilities in different cell types and even in separate compartments of the same cell [22]. Based on this, a multi-tubulin hypothesis was proposed, which stated that in order to perform different cellular functions, specialized microtubule structures are built using diverse tubulin isotypes [23]. This theory was further supported when multiple genes encoding α- and β-tubulin isotypes were identified on distinct chromosomes [24]. So far, seven α-tubulin and nine β-tubulin isotypes have been reported to exist in humans (Table 1). These tubulin isotypes share a high degree of sequence similarity [20]. An overlay of the modeled tubulin isotypes shows that the globular domain of both the α- and β-tubulin isotypes is highly conserved (Figure 3A,B) and the deviations in the sequences are concentrated at the unstructured C-terminal tail, especially in the last 15–20 amino acids (Figure 3C). These variations in the C-terminal tail confer unique qualities to the isotype, allowing it to be differentially modified and regulated by different microtubule interacting proteins [25].

Tubulin isotypes exhibit overall structural homology but diverse C-terminal tails.

Figure 3.
Tubulin isotypes exhibit overall structural homology but diverse C-terminal tails.

Tertiary structures of human α- and β-tubulin isotypes were generated by Modeller v9.17, using 5LYJ as the template structure. The generated structures were then aligned using PyMOL v1.7.4 to show their structural homology. The structural alignment of all the (A) α-tubulin isotypes, TubA1A-blue, TubA1B-cyan, TubA1C-green, TubA3C-lemon green, TubA3E-yellow, TubA4A-orange, TubA8-red and (B) β-tubulin isotypes, TubB1-blue, TubB2A-cyan, TubB2B-green, TubB3-lemon green, TubB4A-yellow, TubB4B-yellow-orange, TubB5-orange, TubB6-salmon red, TubB8-red are shown in cartoon representation. (C) The amino acid sequences of the C-terminal tails of tubulin isotypes indicate sequence diversity. Residues that are not common in all the isotypes are shown in red which are mostly concentrated at the last 15–20 amino acids. The sequences were taken from the NCBI database.

Figure 3.
Tubulin isotypes exhibit overall structural homology but diverse C-terminal tails.

Tertiary structures of human α- and β-tubulin isotypes were generated by Modeller v9.17, using 5LYJ as the template structure. The generated structures were then aligned using PyMOL v1.7.4 to show their structural homology. The structural alignment of all the (A) α-tubulin isotypes, TubA1A-blue, TubA1B-cyan, TubA1C-green, TubA3C-lemon green, TubA3E-yellow, TubA4A-orange, TubA8-red and (B) β-tubulin isotypes, TubB1-blue, TubB2A-cyan, TubB2B-green, TubB3-lemon green, TubB4A-yellow, TubB4B-yellow-orange, TubB5-orange, TubB6-salmon red, TubB8-red are shown in cartoon representation. (C) The amino acid sequences of the C-terminal tails of tubulin isotypes indicate sequence diversity. Residues that are not common in all the isotypes are shown in red which are mostly concentrated at the last 15–20 amino acids. The sequences were taken from the NCBI database.

Table 1
Human tubulin isotypes and their tissue-specific expression
Isotype Gene name NCBI accession no. Tissue-specific expression Reference 
α-tubulin isotypes 
αIa, αIb, αIc TUBA1A/B/C NM_006009.3
NM_006082.2
NM_032704.4 
Widely distributed in all the tissue types, αIa highly expressed in the fetal brain [30
αIIIc TUBA3C P0DPH7 Testis [31
αIIIe TUBA3E NM_207312.2 Testis [32,33
αIVa TUBA4A NM_006000.2 Ubiquitously expressed in all tissue types [32,33
αVIII TUBA8 NM_018943.2 Majorly found in the heart and skeletal muscles and in low levels in the testis and brain [34,35
β-tubulin isotypes 
βI TUBB NM_178014.3 Highly expressed in the spleen, thymus and in the immature brain [29
βIIa, βIIb TUBB2A/2B NM_001069.2
NM_178012.4 
Mainly in the nervous system and in high amounts in neurons and glia [29
βIII TUBB3 NM_006086.3 Neurons of the central and peripheral nervous system, sertoli cells of the testis and in spermatozoa [29,37
βIVa TUBB4 NM_006087.3 The nervous system, especially in neuron, glia, and oligodendrocytes of the brain
Moderate amounts in testis and very low amounts in other tissues 
[28,29
βIVb TUBB2C NM_006088.5 Ciliated cells mostly concentrated in the cilia [29
βV TUBB6 NM_032525.2 Breast and lung tissue [29
βVI TUBB1 NM_030773.3 Hematopoietic tissue, especially leukocytes [29
βVIII TUBB8 NM_177987.2 Oocytes and early embryo [36
Isotype Gene name NCBI accession no. Tissue-specific expression Reference 
α-tubulin isotypes 
αIa, αIb, αIc TUBA1A/B/C NM_006009.3
NM_006082.2
NM_032704.4 
Widely distributed in all the tissue types, αIa highly expressed in the fetal brain [30
αIIIc TUBA3C P0DPH7 Testis [31
αIIIe TUBA3E NM_207312.2 Testis [32,33
αIVa TUBA4A NM_006000.2 Ubiquitously expressed in all tissue types [32,33
αVIII TUBA8 NM_018943.2 Majorly found in the heart and skeletal muscles and in low levels in the testis and brain [34,35
β-tubulin isotypes 
βI TUBB NM_178014.3 Highly expressed in the spleen, thymus and in the immature brain [29
βIIa, βIIb TUBB2A/2B NM_001069.2
NM_178012.4 
Mainly in the nervous system and in high amounts in neurons and glia [29
βIII TUBB3 NM_006086.3 Neurons of the central and peripheral nervous system, sertoli cells of the testis and in spermatozoa [29,37
βIVa TUBB4 NM_006087.3 The nervous system, especially in neuron, glia, and oligodendrocytes of the brain
Moderate amounts in testis and very low amounts in other tissues 
[28,29
βIVb TUBB2C NM_006088.5 Ciliated cells mostly concentrated in the cilia [29
βV TUBB6 NM_032525.2 Breast and lung tissue [29
βVI TUBB1 NM_030773.3 Hematopoietic tissue, especially leukocytes [29
βVIII TUBB8 NM_177987.2 Oocytes and early embryo [36

The tubulin isotypes in humans are differentially expressed in different tissues and are associated with specialized functions [2636]. The tissue-specific expression of all the human tubulin isotypes, along with their gene names and NCBI accession codes, is provided in Table 1. Some of the tubulin isotypes, such as αI and αIVa, are found to be ubiquitously expressed, whereas some of the isotypes are restricted in specialized cells such as βIII in neurons and spermatozoa, βIVb in ciliated and flagellated cells and βVI in blood cells [29,37]. Different tubulin isotypes have been shown to intermingle freely and to participate in microtubule assembly [38]. However, studies in Drosophila and mouse have noted that substitution of one isotype by another leads to a loss of the specialized function implying that the isotypes are functionally non-interchangeable [39,40]. This suggests that tubulin isoforms may regulate the microtubule dynamics by regulating the intrinsic polymer properties.

Post-translational modifications of tubulin

Apart from tubulin isotypes, PTMs of tubulins also contribute to tubulin heterogeneity [41]. Both α- and β-tubulins are subjected to a wide range of PTMs (Figure 4A and Table 2). Most of the tubulin PTMs, such as tyrosination–detyrosination, (poly) glutamylation, (poly) glycylation, acetylation, phosphorylation, and methylation, are well studied [42]. However, PTMs, such as polyamination, palmitoylation, ubiquitination, glycosylation, arginylation, sumoylation, succination, and O-Glc-NAcylation, have been reported to occur on tubulin but are yet to be investigated in detail [43]. The PTMs of tubulins have been reviewed earlier [4244] and hence will not be elaborated here. However, important PTMs with their modifying enzymes and type of microtubules on which they commonly occur are summarized in Table 2.

Post-translational modifications (PTMs) of tubulin and their localization patterns in different cells.

Figure 4.
Post-translational modifications (PTMs) of tubulin and their localization patterns in different cells.

(A) A tubulin heterodimer (PDB ID: 5LYJ) is shown in surface representation with α-tubulin in green and β-tubulin in blue. Tubulin PTMs occurring on specific sites of the heterodimer are shown. The amino acids of the C-terminal tails are denoted by a single letter code and the modification sites for polyglycylation and polyglutamylation shown are randomly chosen. Am, amination; Ph, phosphorylation; Ac, acetylation; Me, methylation, Δ2-tubulin and Δ3-tubulin are α-tubulin without two (EY) and three (EEY) extreme C-terminal residues, respectively. (B) The diverse patterns of PTMs on various microtubule structures in different cells are shown. The cells are as mentioned in Figure 1. The microtubule arrangement in the axoneme of non-motile primary cilia, motile cilia and centrioles are shown in cyan, orange and red, respectively. The microtubules of the marginal band in the platelets are also shown in blue and purple.

Figure 4.
Post-translational modifications (PTMs) of tubulin and their localization patterns in different cells.

(A) A tubulin heterodimer (PDB ID: 5LYJ) is shown in surface representation with α-tubulin in green and β-tubulin in blue. Tubulin PTMs occurring on specific sites of the heterodimer are shown. The amino acids of the C-terminal tails are denoted by a single letter code and the modification sites for polyglycylation and polyglutamylation shown are randomly chosen. Am, amination; Ph, phosphorylation; Ac, acetylation; Me, methylation, Δ2-tubulin and Δ3-tubulin are α-tubulin without two (EY) and three (EEY) extreme C-terminal residues, respectively. (B) The diverse patterns of PTMs on various microtubule structures in different cells are shown. The cells are as mentioned in Figure 1. The microtubule arrangement in the axoneme of non-motile primary cilia, motile cilia and centrioles are shown in cyan, orange and red, respectively. The microtubules of the marginal band in the platelets are also shown in blue and purple.

Table 2
Tubulin post-translational modifications
PTM Modification The site on α- or β-tubulin Forward enzyme Reverse enzyme Cellular MT 
Acetylation Acetyl group addition Lys40 of α-tubulin αTAT1 HDAC6 and SirT2 Axonemal MT, spindle, midbody and marginal band MTs 
Lys252 of β-tubulin San acetyltransferase 
Detyrosination C-terminal tyrosine removal Terminal tyrosine on C-terminal of α-tubulin Vasohibin family member TTL Kinetochore fibers, midbody, axonemal MT and marginal band MTs 
Tyrosination Addition of tyrosine C-terminal of α-tubulin TTL Vasohibin family member Interphase MT and growth cone MT 
Δ2- and Δ3-tubulin Removal of C-terminal glutamate from detyrosinated α-tubulin CTT of α-tubulin CCP family Not reported Axonal MT 
Glutamylation Addition of one or more glutamates as branched peptide chains Multiple glutamates on the α- and β-tubulin CTT TTLL1, 4–7, 9, 11, 13 CCP deglutamylase Kinetochore fibers, midbody, and axonemal MT 
Glycylation Addition of one or more glycines as branched peptide chains Multiple glutamates on the α- and β-tubulin CTT TTLL3, 8, 10 Not reported Axonemal MT 
Phosphorylation Addition of phosphate Ser172 of β-tubulin Cyclin-dependent kinase 1 (Cdk1) Not reported Unknown 
Unidentified tyrosine on CTT of α-tubulin Kinase Syk 
Polyamination Addition of polyamines Glutamines of α- and β-tubulin Unknown transglutaminase Not reported Axonemal MT 
Methylation Addition of a methyl group Lys40 of α-tubulin SETD2 methyltransferase Not reported Central spindle MT (not astral) 
PTM Modification The site on α- or β-tubulin Forward enzyme Reverse enzyme Cellular MT 
Acetylation Acetyl group addition Lys40 of α-tubulin αTAT1 HDAC6 and SirT2 Axonemal MT, spindle, midbody and marginal band MTs 
Lys252 of β-tubulin San acetyltransferase 
Detyrosination C-terminal tyrosine removal Terminal tyrosine on C-terminal of α-tubulin Vasohibin family member TTL Kinetochore fibers, midbody, axonemal MT and marginal band MTs 
Tyrosination Addition of tyrosine C-terminal of α-tubulin TTL Vasohibin family member Interphase MT and growth cone MT 
Δ2- and Δ3-tubulin Removal of C-terminal glutamate from detyrosinated α-tubulin CTT of α-tubulin CCP family Not reported Axonal MT 
Glutamylation Addition of one or more glutamates as branched peptide chains Multiple glutamates on the α- and β-tubulin CTT TTLL1, 4–7, 9, 11, 13 CCP deglutamylase Kinetochore fibers, midbody, and axonemal MT 
Glycylation Addition of one or more glycines as branched peptide chains Multiple glutamates on the α- and β-tubulin CTT TTLL3, 8, 10 Not reported Axonemal MT 
Phosphorylation Addition of phosphate Ser172 of β-tubulin Cyclin-dependent kinase 1 (Cdk1) Not reported Unknown 
Unidentified tyrosine on CTT of α-tubulin Kinase Syk 
Polyamination Addition of polyamines Glutamines of α- and β-tubulin Unknown transglutaminase Not reported Axonemal MT 
Methylation Addition of a methyl group Lys40 of α-tubulin SETD2 methyltransferase Not reported Central spindle MT (not astral) 

Abbreviations: MT, microtubule; CTT, C-terminal tail; CCP, cytoplasmic carboxypeptidase; TTL, tubulin tyrosine ligase; TTLL, tubulin tyrosine ligase-like.

Although the majority of the PTMs take place on the C-terminal tails of the α- and β-tubulins, few PTMs, such as acetylation [45], phosphorylation [46], polyamination [47] and methylation [48], occur on the structured core of tubulin (Figure 4A). Among these, acetylation of α-tubulin is a unique modification, as it occurs on lysine 40, which is present in the lumen of the cylindrical microtubules [45]. Recently, αTAT1, a tubulin acetyltransferase, was shown to enter the microtubule lumen either through the ends of the microtubules or through the breaks and bends in the microtubule lattice [49]. Most of the PTMs occur on tubulin heterodimers that are already polymerized into microtubules. However, some modifications, such as phosphorylation and polyamination, take place on free soluble tubulin, which when polymerized can alter the intrinsic polymer properties [46,47]. Specific patterns of PTMs can program the microtubules to perform specialized functions, either by modulating the mechanical properties of microtubules or by influencing the interaction of microtubules with MAPs [5053]. These patterns vary in different cell types, intracellular region and even along the same microtubule (Figure 4B). Moreover, overall patterns of the PTMs also depend upon the presence of different tubulin isotypes, thereby adding an additional layer of complexity to the regulatory mechanism.

The tubulin code and its influence on the microtubule dynamics

The diversity in microtubule structure arising from the incorporation of multiple tubulin isotypes and their numerous PTMs gives rise to a complex signaling mechanism known as the ‘tubulin code’ [44,54]. The tubulin code provides a guidance system for crucial cellular processes, such as intracellular signaling, targeted vesicle transport, and cell division and differentiation [55]. The tubulin code generates unique signals by combining the various tubulin isotypes and their PTMs. These signals are then read by microtubule effectors, such as MAPs, +TIPs and motor proteins, which alter the microtubule dynamics based on the signal [54]. The complex signal arising from the code can also modulate the interaction of MAPs and motors with microtubules.

Influence of tubulin isotypes on microtubule dynamics

Microtubules composed of different tubulin isotypes exhibit differential dynamic properties [56]. Specifically, microtubules composed of the βIII-tubulin isotypes exhibit more dynamicity than the microtubules formed of the βII or βIV isotypes, indicating that different compositions of tubulin isotypes can alter the dynamicity of microtubules [56]. In a recent study, microtubules when assembled from tubulin purified from human embryonic kidney (HEK) cell lines, which contain mostly the βI and βIVb isotypes, exhibited a faster growth rate and low catastrophe frequency than microtubules assembled from brain tubulin [57]. Moreover, the dynamics of the non-neuronal (HEK cells) microtubules changed when neuronal (brain) tubulin was added, indicating that the cells can tune microtubule dynamics by varying the relative expression level of certain tubulin isotypes [57]. In addition to its role in the regulation of microtubule stability, tubulin isotypes have also been reported to regulate the protofilament number of microtubules [58]. Microtubules composed of isotypically purified human βIIb tubulin exhibited a broad distribution of the protofilament number and were more stable than the microtubules composed of βIII-tubulin isotypes which had canonical 13 protofilaments [58]. Specialized microtubules, such as axonemal, neuronal and marginal band microtubules, are also specifically enriched with particular β-tubulin isotypes [27,59,60]. Considering the special role of such microtubules in cells, any alteration in the expression levels of the tubulin isotypes can alter the overall microtubule function. The incorporation of different tubulin isotypes into the protofilament can occur for two primary reasons: (1) Subtle alterations in the structured core of the tubulin isotypes can influence the physical properties and assembly kinetics of the microtubules, thus affecting the stability of microtubules. (2) Variations in the sequence of the C-terminal tail, a primary binding site for several MAPs, can differentially regulate the interactions of MAPs and microtubules. Similarly, mutations in the tubulin genes can also alter tubulin structure and hence affect the functions of the microtubules.

Influence of tubulin PTMs on microtubule dynamics

PTMs constitute the second component of the tubulin code. PTMs that occur on the structured core of tubulin can disturb its conserved fold, thereby perturbing the microtubule lattice and overall behavior and functions of the microtubules [50,51,61]. For instance, acetylation that occurs in the conserved region of the tubulin core increases the mechanical resilience of the microtubule lattice by reducing lateral contacts between the protofilaments [50]. This makes the microtubules resistant to mechanical stress and breakage (Figure 5A, i), thereby explaining the long life of most of the acetylated microtubules [62]. In contrast, a large number of modifications occur on the genetically diverse C-terminal tails of tubulin, where different patterns of PTMs influence microtubule dynamics [43]. For example, stable long-lived microtubules are generally detyrosinated, whereas tyrosination is a hallmark of highly dynamic microtubules (Figure 5A, ii,iii) [43]. Polyglutamylation has also been shown to regulate the dynamicity of microtubules by controlling the microtubule severing activity of spastin, an ATPase (Figure 5B, i). Spastin recognizes polyglutamated microtubules and responds to the number of glutamates added to tubulin [63].

Effect of tubulin PTMs on microtubule stability and interaction of motor proteins.

Figure 5.
Effect of tubulin PTMs on microtubule stability and interaction of motor proteins.

Microtubules with specific PTMs are shown. Unmodified microtubules (without any modification), acetylated microtubules (acetyl group is indicated by an orange star), detyrosinated microtubules (terminal tyrosine residue ‘Y’ absent at the C-terminal tail of α-tubulin), tyrosinated microtubules (terminal tyrosine residue ‘Y’ present at the C-terminal tail of α-tubulin), and polyglutamylated microtubules (branched C-terminal tail). (A) Effects of tubulin PTMs on the stability of microtubules. (i) Under mechanical stress, straight unmodified microtubules undergo breakage due to the bending of microtubules. However, acetylated microtubules can resist the breakage due to enhanced mechanical resilience. (ii) Acetylated and detyrosinated microtubules are stable, whereas (iii) tyrosinated microtubules are dynamic. (B) Effects of tubulin PTMs on the interaction of motor and microtubule-associated proteins. The association of motors and MAPs with differentially modified microtubules is shown. (i) Spastin identifies polyglutamylated microtubules and causes microtubule severing. Axonemal dynein preferentially interacts with polyglutamylated microtubules of the axonemes. (ii) CENP-E selectively interacts with detyrosinated kinetochore microtubules to guide chromosomes to the metaphase plate. Kinesin-1 binds to detyrosinated stable microtubules and assists in axonal transport. (iii) Kinesin-13 induces microtubule depolymerization by preferentially binding to tyrosinated microtubules. The dynein–dynactin complex associates with tyrosinated microtubules for the initiation of dynein-driven motility.

Figure 5.
Effect of tubulin PTMs on microtubule stability and interaction of motor proteins.

Microtubules with specific PTMs are shown. Unmodified microtubules (without any modification), acetylated microtubules (acetyl group is indicated by an orange star), detyrosinated microtubules (terminal tyrosine residue ‘Y’ absent at the C-terminal tail of α-tubulin), tyrosinated microtubules (terminal tyrosine residue ‘Y’ present at the C-terminal tail of α-tubulin), and polyglutamylated microtubules (branched C-terminal tail). (A) Effects of tubulin PTMs on the stability of microtubules. (i) Under mechanical stress, straight unmodified microtubules undergo breakage due to the bending of microtubules. However, acetylated microtubules can resist the breakage due to enhanced mechanical resilience. (ii) Acetylated and detyrosinated microtubules are stable, whereas (iii) tyrosinated microtubules are dynamic. (B) Effects of tubulin PTMs on the interaction of motor and microtubule-associated proteins. The association of motors and MAPs with differentially modified microtubules is shown. (i) Spastin identifies polyglutamylated microtubules and causes microtubule severing. Axonemal dynein preferentially interacts with polyglutamylated microtubules of the axonemes. (ii) CENP-E selectively interacts with detyrosinated kinetochore microtubules to guide chromosomes to the metaphase plate. Kinesin-1 binds to detyrosinated stable microtubules and assists in axonal transport. (iii) Kinesin-13 induces microtubule depolymerization by preferentially binding to tyrosinated microtubules. The dynein–dynactin complex associates with tyrosinated microtubules for the initiation of dynein-driven motility.

Influence of tubulin PTMs on microtubule motors

Tubulin PTMs can also influence the microtubule dynamics by regulating the interaction of motor proteins with microtubules (Figure 5B) [6467]. Polyglutamylation, found abundantly on axonemes (Figure 4B), has been shown to modulate the activity of ciliary dynein motors, thereby controlling the beating of cilia (Figure 5B, i) [64,65,68]. Distinct PTM signatures can create subpopulations of microtubules that differ in their dynamic properties and hence are recognized by a different set of motors for differential cargo transport [52,53]. For example, the kinetochore microtubules of the mitotic spindle are specifically detyrosinated, whereas the astral microtubules are not (Figure 4B). CENP-E, a plus-end-directed motor, has been shown to have enhanced motility on detyrosinated microtubules (Figure 5B, ii) [52]. This allows the CENP-E motor to preferentially bind to kinetochore microtubules and guide the chromosome towards the metaphase plate [52]. In neurons, different subpopulations of microtubules exist to control the overall cargo transport. The microtubules of axons and dendrites are differentially modified as microtubules of the dendrites are tyrosinated, whereas those of axons are detyrosinated (Figure 4B) [53]. These two subsets of microtubules are utilized by different sets of motors that transport different cargos to distinct locations [53,69]. For example, kinesin-1, a plus-end-directed motor, has been shown to preferentially bind to detyrosinated microtubules and assist in cargo transport to the axons (Figure 5B, ii) [70,71]. However, using chimeric yeast tubulin, kinesin-1 has been shown to exhibit more processivity on polyglutamylated microtubules [72]. In addition, acetylation has been shown to enhance the activity of kinesin-1 indicating that different PTMs can modulate motor activity [73]. Kinesin-13, a microtubule depolymerizer, has been shown to preferentially bind to tyrosinated microtubules rather than detyrosinated microtubules (Figure 5B, iii) [67]. Detyrosination, observed mostly on long-lived microtubules, enhances microtubule stability by preventing the interaction of kinesin-13 motors [67]. Tyrosination has been reported to be crucial for the initiation of the dynein–dynactin motility (Figure 5B, iii) [74]. The p150 subunit of dynactin interacts with the terminal tyrosine residue of the α-tubulin C-terminal tail and initiates dynein-driven motility [74].

Influence of the tubulin code on cellular processes

Several cellular processes that are essential for the growth and development of cells are regulated by distinct signatures generated by the tubulin code [44]. During cell differentiation, many changes occur in the expression levels of certain tubulin PTMs, which alter the proportion of stable and dynamic microtubules in the cell. For instance, the fine-tuning of polyglutamylation is critical during postnatal development [75]. During the initial stages, extensive polyglutamylation is required for cerebellar development, but a strict reduction in its levels at the end of the neuron growth is equally important. A perturbation in the tuning of glutamylation leads to detrimental effects on the development of the cell [75]. The detyrosination–tyrosination cycle of α-tubulin has also been recognized to be crucial for cell cycle progression and cell differentiation as it plays a significant role in organelle and vesicular transport [76,77]. The cellular role of glycylation has not been comprehensively studied, but it is reported to be mostly found on cilia where it maintains the integrity of the motile cilia. It is also involved in regulating the length of the non-motile primary cilia (Figure 4B) [66]. Specific PTMs that occur on particular isotypes of tubulin have also been shown to selectively regulate the velocity and processivity of certain microtubule motors [72]. Thus, tubulin isotypes and their PTMs can together generate many possible combinations of complex signals that have the potential to regulate the microtubule dynamicity by controlling the recruitment and behavior of microtubule regulators.

Tubulin heterogeneity and cancer

Owing to several important roles in cell division, microtubules have been considered as excellent targets for anticancer agents. Over the years, several microtubule-targeting agents (MTAs) or tubulin-binding agents (TBAs) have been employed as chemotherapeutic agents [78]. TBAs directly bind to either tubulin dimers or microtubules and perturb the dynamicity of microtubules. Based on their effects on microtubules, TBAs are divided into two groups, namely, microtubule-polymerizing/stabilizing agents and microtubule-depolymerizing agents. TBAs that promote the polymerization of microtubules are paclitaxel, docetaxel, and epothilones, whereas Vinca alkaloids, dolastatin, combretastatin, and 2-methoxyestradiol are known to depolymerize microtubules [79,80]. However, the majority of the TBAs inhibit cancer cell proliferation by suppressing the dynamics of microtubules without significantly altering the level of polymerized tubulin [78]. As TBAs suppress spindle microtubule dynamics, the attachment of microtubules to kinetochores is disturbed, thereby hampering chromosome congression. A delay in chromosome congression halts the metaphase to anaphase transition and arrests cells at mitosis by activating the spindle assembly checkpoint [78]. A prolonged mitotic block induces apoptosis, thereby causing cell death. Hence, targeting microtubules of highly proliferating cancerous cells is an interesting approach for cancer treatment.

Alterations in tubulin isotype expression

The change in the expression levels of tubulin isotypes has been extensively studied in a variety of tumors including solid and hematological cancers (Table 3). In most of the cancer types, expressions of certain tubulin isotypes are found to be up-regulated [8185]. Also, the extent of the increase in the isotype expression level corresponds to the aggressive form of the condition and poor patient outcome [83,86,87]. Among the different β-tubulin isotypes, abnormal expression of βIII-tubulin has been detected in a wide range of tumors such as lung, breast, ovarian, prostate, uterine and gastric cancers (Table 3) [8892]. Recently, βIII was also found to be highly expressed in most of the head and neck squamous cell cancers [93]. Apart from βIII, differential expression of βI, βII, and βIV has been detected in breast, lung and ovarian cancers (Table 3) [9497]. Another isotype, βV, which is generally not expressed in fallopian tube epithelium, has been found to be up-regulated in serous ovarian neoplasms [98] and also correlates with high-grade serous carcinomas. Aberrant expression of βV has also been detected in breast and lung cancers [99]. Although altered expressions of several β-tubulin isotypes have been reported, their relevance in terms of clinical studies still requires extensive investigation. However, several clinical and in vitro studies have been performed to understand the role of an increased βIII level in the development of chemoresistance (Table 3) [100].

Table 3
Alterations in tubulin isotype expression and its significance in cancer
Isotype Alteration Cancer Outcome Reference 
αIb-tubulin Increased expression Hepatocellular carcinoma Resistance to paclitaxel and poor prognosis [81
Mantle cell lymphoma Poor prognosis [86
βI-tubulin Increased expression Ovarian cancer Resistance to taxol [83
Breast cancer Weak response to docetaxel treatment [94
βII-tubulin Increased expression LASCCHN Poor survival rate when treated with docetaxel [87
Decreased expression Ovarian and breast cancer Resistance to taxane treatment and correlates with advanced stage of the condition [95,96
Depletion NSCLC cell line Increased sensitivity to vinca alkaloids [101
βIII-tubulin Increased expression levels Ovarian cancer Resistance to paclitaxel that correlates with poor clinical outcome [83
Breast cancer Weak response to taxane treatment [82,88
NSCLC Resistance to taxane [89
Gastric cancer Weak response to taxane treatment [90
Prostate Resistance to docetaxel [91
Uterine serous carcinoma Weak response to taxane treatment [92
Depletion NSCLC cell line Increased sensitivity towards docetaxel and cisplatin, and towards epothilone B [89,104
PDA cell line Enhanced chemosensitivity [102
βIVa-tubulin Increased expression Ovarian cancer Weak response to taxol treatment [83
βIVb-tubulin Decreased expression Breast cancer cell line Resistance to docetaxel [97
Depletion Lung cancer cell line and PDA cell line Increased sensitivity to vinca alkaloids [101,102
NSCLC cell line Increased sensitivity to vinca alkaloids [101
βV-tubulin Increased expression NSCLC A positive response to taxanes [84,85
Serous ovarian neoplasms Expression level correlates with the grade of the carcinoma [98
Isotype Alteration Cancer Outcome Reference 
αIb-tubulin Increased expression Hepatocellular carcinoma Resistance to paclitaxel and poor prognosis [81
Mantle cell lymphoma Poor prognosis [86
βI-tubulin Increased expression Ovarian cancer Resistance to taxol [83
Breast cancer Weak response to docetaxel treatment [94
βII-tubulin Increased expression LASCCHN Poor survival rate when treated with docetaxel [87
Decreased expression Ovarian and breast cancer Resistance to taxane treatment and correlates with advanced stage of the condition [95,96
Depletion NSCLC cell line Increased sensitivity to vinca alkaloids [101
βIII-tubulin Increased expression levels Ovarian cancer Resistance to paclitaxel that correlates with poor clinical outcome [83
Breast cancer Weak response to taxane treatment [82,88
NSCLC Resistance to taxane [89
Gastric cancer Weak response to taxane treatment [90
Prostate Resistance to docetaxel [91
Uterine serous carcinoma Weak response to taxane treatment [92
Depletion NSCLC cell line Increased sensitivity towards docetaxel and cisplatin, and towards epothilone B [89,104
PDA cell line Enhanced chemosensitivity [102
βIVa-tubulin Increased expression Ovarian cancer Weak response to taxol treatment [83
βIVb-tubulin Decreased expression Breast cancer cell line Resistance to docetaxel [97
Depletion Lung cancer cell line and PDA cell line Increased sensitivity to vinca alkaloids [101,102
NSCLC cell line Increased sensitivity to vinca alkaloids [101
βV-tubulin Increased expression NSCLC A positive response to taxanes [84,85
Serous ovarian neoplasms Expression level correlates with the grade of the carcinoma [98

Abbreviations: LASCCHN, locally advanced squamous cell cancer of the head and neck; NSCLC, non-small cell lung cancer; PDA, pancreatic ductal adenocarcinoma.

Altered tubulin isotype levels confer resistance to TBAs

Several studies carried out in different types of tumors have noted that increased expression of β-tubulin isotypes imparts resistance against anti-tubulin agents (Table 3). Elevated levels of βI, βII, βIII, βIV and βV have been reported to confer resistance to taxanes in a wide range of tumor types (Table 3). Several in vitro studies are also in agreement with these observations, as several drug-resistant cancer cell lines exhibit an altered expression profile of β-tubulin isotypes (Table 3) [89,97,101,102]. Among the different β-tubulin isotypes, the role of βIII-tubulin in the development of resistance has been extensively studied. Elevated βIII levels have been observed to confer resistance mainly against taxanes in a wide variety of cancers (Table 3). Moreover, a positive correlation between βIII expression level and the advanced stage of ovarian cancer has been established. As a severe form of the condition exhibits an enhanced βIII level and correspondingly poor clinical outcome [83,100]. The overexpression of the βIII isotype in various cancers and its correlation with drug resistance suggest that βIII isotype opposes the suppressing effects of TBAs on microtubule dynamics. Microtubules formed from purified αβIII isotype are more dynamic than the microtubules formed from either αβII or αβIV isotype [56]. Also, microtubules consisting of αβIII-tubulin exhibits a 7.4-fold less sensitivity to paclitaxel than the microtubules formed from a mixture of tubulin isotypes [103]. This indicates that an increased level of βIII-tubulin confers resistance against taxanes by inhibiting the ability of taxanes to stabilize microtubules. Consistent with this, the depletion of βIII-tubulin resulted in increased chemosensitivity of cancer cells towards a broad range of chemotherapy drugs [102,104]. Several studies also reported a correlation between increased βIII level and poor patient outcome [82,83,88,90]. Therefore, it is hypothesized that an increase in the level of βIII may assist cells to thrive under stressed conditions, which ultimately leads to cell survival. In a recent study, a colchicine-resistant breast cancer cell line was shown to exhibit a 2.3-fold increase in the level of βIII-tubulin [105]. It was speculated that the enhanced βIII expression conferred resistance against colchicine. Interestingly, microtubule dynamics were suppressed in these cells, and the cells were more sensitive to paclitaxel [105]. A few studies have also reported contradictory findings where elevated βIII level failed to confer resistance to certain TBAs [106108]. This suggests that the altered βIII expression modulates the sensitivity of cells to different TBAs, in an unequal manner.

Alteration in tubulin PTMs

Alterations in the PTMs of microtubules have been observed in a wide range of cancers (Table 4). The regulation of the levels of tubulin PTMs occurs on multiple levels including the regulation of the expression level of the modifying enzymes that bring about a specific PTM. The division machinery of the cell, such as meiotic or mitotic spindle, midbody, and centrioles, possesses a considerable number of tubulin PTMs that impart a precise control over the cell division [55]. For example, a perturbation in the detyrosination of kinetochore microtubules can hamper the transport of chromosomes, leading to aneuploidy [52]. A disturbance in the detyrosination–tyrosination cycle has been reported to affect the progression of a wide range of cancers (Table 4) [109112]. In a survey of human tumor biopsies, tumors of different tissue origins displayed loss of tubulin tyrosine ligase (TTL) activity, which was speculated to enhance the growth of the tumors [109]. Furthermore, the down-regulation of TTL activity in breast cancer cells has been reported to induce epithelial-to-mesenchymal transition (EMT), which increases the metastatic potential of the cells [113]. The suppression of TTL activity and an imbalanced carboxypeptidase (CCP) activity together provide a strong selective advantage for the survival of cancer cells. Recently, the discovery of vasohibins as detyrosination-catalyzing enzymes has opened up several new links between previously known vasohibin dysfunctions and cancer [114,115]. An increase in tubulin acetylation levels has also been associated with cancer cells as acetylation is involved in the maintenance of cellular homeostasis [116]. Increased HDAC6 activity has been identified as a marker for better prognosis in breast cancer where patients with elevated HDAC6 levels responded better to endocrine treatment than did those with low levels [117]. A study in breast cancer cells also showed that pharmacological inhibition of HDAC6 increases its binding to microtubules, thereby suppressing the dynamic instability of the microtubules [118]. Other tubulin modifications have also been linked to cancer. An increased level of polyglutamylation has been observed in breast cancer where it confers resistance to paclitaxel (Table 4) [119]. A decrease in tubulin glycylation levels has been reported in colorectal cancers [120]. Δ2-tubulin modification of the βIVb isotype has been observed in the advanced stages of liver cancer in humans and mouse models of hepatic carcinoma [121]. All these alterations in the patterns of the microtubule PTMs more likely perturb the homeostasis of the cancer cells; however, the mechanism by which they assist the cancer cells is not yet known. Spatiotemporal mapping of the tubulin PTMs may help to elucidate the role of the tubulin PTMs in intracellular signaling and cell survival.

Table 4
Alterations in tubulin PTMs in cancer
PTM Alteration Cancer Outcome Reference 
Tyrosination–detyrosination Suppression of TTL activity Non-epithelial tumors of different origins Loss of TTL activity enhances tumor growth [109
Down-regulated TTL expression Prostate cancer cell line Elevated levels of detyrosinated and polyglutamylated α-tubulin [110
Breast cancer cells Linked to tumor aggressiveness and promotes metastasis [111,113
Differential TTL expression Neuroblastoma Correlates with poor prognosis and patient outcome [112
Acetylation Elevated HDAC6 levels Breast cancer A marker for better prognosis [117
Glutamylation Increased levels of polyglutamylated tubulins Breast cancer cells Paclitaxel-resistant [119
Glycylation Down-regulation of TTLL3 Colorectal cancers Increased rate of cell proliferation due to loss of primary cilia [120
Δ2-tubulin Increased Δ2-tubulin modification of βIVb Hepatic carcinoma Corresponds to advanced stages of liver cancer [121
PTM Alteration Cancer Outcome Reference 
Tyrosination–detyrosination Suppression of TTL activity Non-epithelial tumors of different origins Loss of TTL activity enhances tumor growth [109
Down-regulated TTL expression Prostate cancer cell line Elevated levels of detyrosinated and polyglutamylated α-tubulin [110
Breast cancer cells Linked to tumor aggressiveness and promotes metastasis [111,113
Differential TTL expression Neuroblastoma Correlates with poor prognosis and patient outcome [112
Acetylation Elevated HDAC6 levels Breast cancer A marker for better prognosis [117
Glutamylation Increased levels of polyglutamylated tubulins Breast cancer cells Paclitaxel-resistant [119
Glycylation Down-regulation of TTLL3 Colorectal cancers Increased rate of cell proliferation due to loss of primary cilia [120
Δ2-tubulin Increased Δ2-tubulin modification of βIVb Hepatic carcinoma Corresponds to advanced stages of liver cancer [121

Current advances and future perspective

Development of potent anti-tubulin agents to overcome isotype-specific resistance

Currently, a large number of TBAs are being used as clinical agents against different types of cancers [122]. However, the development of resistance against the TBAs is an enormous obstacle for cancer treatment. Therefore, persistent efforts are being made to improve the potency of drugs against resistant cell lines and to increase their tumor specificity. Several TBAs such as noscapine [123], laulimalide [124], peloruside [125], taccalonolide [126] and zampanolide [127] have been shown to potently inhibit the proliferation of taxane-resistant cancer cells. Novel TBAs are considered to be potent based on their ability to be effective against drug-resistant cells. In addition, agents that show insensitivity towards increased expression of tubulin isotype particularly, βIII-tubulin, have been selected for further clinical studies [128,129]. An increase in βIII expression level has been suggested as a cause of development of taxane resistance in many cancer cells [83,89,90,92]. Therefore, the inhibitors that are sensitive to such elevated βIII levels are preferred for countering taxane resistance. Ixabepilone, an analog of epothilone B, has been reported to be active in taxane-resistant metastatic breast cancer cells [130]. Ixabepilone preferentially suppresses βIII-tubulin dynamics and was found to be more potent against cells expressing elevated βIII-tubulin [130]. Cabazitaxel, a novel taxane, was reported to be more potent against βIII-tubulin expressing breast cancer cells and purified βIII-tubulin-enriched microtubules thus indicating its enhanced anti-tumor efficacy in βIII-tubulin overexpressing tumors [131,132]. Recently, vinblastine analogs were shown to be more potent against increased βIII-tubulin expression in taxol-resistant NSCLC cells [133]. Similarly, DJ101, which targets the colchicine site of tubulin, inhibited tumor growth in a paclitaxel-resistant xenograft mouse model of human prostate cancer indicating that it could be further developed to be used in cancer therapy [134]. Also, a novel quinolone chalcone-based compound that binds at the colchicine site was found to be highly effective against multi-drug-resistant cancer cells in which paclitaxel, colchicine, and vinblastine were ineffective. It also exhibited strong anticancer activity in triple negative breast cancer cells [135]. The development of these agents with enhanced potency towards resistant cells provides an interesting approach to deal with chemotherapy resistance.

Understanding the significance of tubulin heterogeneity

In addition to developing potent anti-tubulin agents, it is essential to gain a comprehensive understanding of the cellular functions of the tubulin code. To do that, we need to understand the influence of each tubulin modification on the microtubule functions. Over the years, mammalian brain tissues have been used to isolate tubulin. However, the isolated tubulins are a mixture of various tubulin isotypes and are also heavily post-translationally modified, thereby making them unsuitable for studying the role of tubulin heterogeneity [136,137]. Earlier attempts were made to purify recombinant tubulin in the bacterial expression system [138]. However, these attempts failed because the machinery required for tubulin folding is absent in bacteria [139,140]. As a substitute, endogenous tubulin was isolated from cultured cell lines such as HeLa cells [141,142]. The purified protein was further subjected to modifying enzymes in vitro to generate specifically modified microtubules [143]. Using the above approach, preferential binding of CENP-E motor to detyrosinated microtubules was demonstrated [52]. Tubulin was isolated from HeLa cells and treated with CCP A to generate detyrosinated tubulins, which was polymerized to form detyrosinated microtubules [52]. Similarly, to generate endogenously modified tubulin, cultured cells were transfected with the desirable modifying enzyme [144]. The influence of different types of polyglutamylation on microtubule severing was studied by isolating endogenously polyglutamylated tubulin from HeLa cells transfected with different glutamylating enzymes [145]. Furthermore, an affinity chromatography method was developed using the microtubule-binding TOG domain of yeast as an affinity matrix to purify cytoplasmic tubulin from any cell type or tissue [146]. Although this method yields small amounts of purified tubulin, it is sufficient to study the tissue-specific tubulin composition and distribution [146]. Using this method, tubulin was purified from HEK cells, and its isoform composition and dynamicity were compared with those of brain tubulin [57]. Using the TOG affinity-based method, the role of graded control of tubulin glutamylation in microtubule severing [63] and the impact of the tubulin code on the motility of axonemal dynein motors were investigated [147]. As the yeast system lacks tubulin-modifying enzymes, it has been used as a source to purify unmodified tubulin [148]. Since the core regions of mammalian and yeast tubulin are highly similar, tubulin chimeras consisting of the yeast tubulin core and the mammalian C-terminal tails were prepared [72]. Using such tubulin chimeras, the regulation of motor proteins by different tubulin isotypes and PTMs was studied [72]. Also, the essential role of α-tubulin tyrosination in the initiation of the dynein–dynactin motility was reported [74]. The co-expression of recombinant human tubulin isotypes with affinity tags has also been successfully optimized in insect cell lines, and the proteins are purified through affinity chromatography followed by cleavage of the affinity tag [58,149]. The advances in the field of tubulin purification over the years have made it possible today to purify recombinant tubulin to a certain extent. In addition to obtaining pure tubulin, identification and purification of several unknown tubulin-modifying enzymes are necessary to determine the role of tubulin PTMs in the tubulin code. Understanding the mechanism and functions of all the tubulin PTMs and isotypes will help to decode the tubulin code, which may provide clues to develop more efficacious anti-tumor drugs.

Abbreviations

     
  • CCP

    carboxypeptidase

  •  
  • CENP-E

    centrosome-associated protein E

  •  
  • CTT

    C-terminal tail

  •  
  • EMT

    epithelial-to-mesenchymal transition

  •  
  • HDAC6

    histone deacetylase 6

  •  
  • LASCCHN

    locally advanced squamous cell cancer of the head and neck

  •  
  • MAPs

    microtubule-associated proteins

  •  
  • MT

    microtubule

  •  
  • MTAs

    microtubule-targeting agents

  •  
  • NSCLC

    non-small cell lung cancer

  •  
  • PDA

    pancreatic ductal adenocarcinoma

  •  
  • PTMs

    post-translational modifications

  •  
  • TBAs

    tubulin-binding agents

  •  
  • TOG

    tumor-overexpressed gene

  •  
  • TTL

    tubulin tyrosine ligase

  •  
  • TTLL

    tubulin tyrosine ligase-like

Funding

This work is supported by a grant [BT/PR14618/BRB/10/1418/2015] from the Department of Biotechnology, Government of India to D.P.

Acknowledgement

We thank Dr. Richard Lueduena, University of Texas, San Antonio, Texas for critical reading of the manuscript. SSP thanks Council of Scientific and Industrial Research, Government of India for her fellowship.

Competing Interests

The Authors declare that there are no competing interests associated with the manuscript.

References

References
1
Desai
,
A.
and
Mitchison
,
T.J.
(
1997
)
Microtubule polymerization dynamics
.
Annu. Rev. Cell Dev. Biol.
13
,
83
117
2
Mitchison
,
T.
and
Kirschner
,
M.
(
1984
)
Dynamic instability of microtubule growth
.
Nature
312
,
237
242
3
Cassimeris
,
L.U.
,
Walker
,
R.A.
,
Pryer
,
N.K.
and
Salmon
,
E.D.
(
1987
)
Dynamic instability of microtubules
.
Bioessays
7
,
149
154
4
Nogales
,
E.
,
Whittaker
,
M.
,
Milligan
,
R.A.
and
Downing
,
K.H.
(
1999
)
High-resolution model of the microtubule
.
Cell
96
,
79
88
5
Felgner
,
H.
,
Frank
,
R.
and
Schliwa
,
M.
(
1996
)
Flexural rigidity of microtubules measured with the use of optical tweezers
.
J. Cell Sci.
109
,
509
516
PMID:
[PubMed]
6
Saxton
,
W.M.
,
Stemple
,
D.L.
,
Leslie
,
R.J.
,
Salmon
,
E.D.
,
Zavortink
,
M.
and
McIntosh
,
J.R.
(
1984
)
Tubulin dynamics in cultured mammalian cells
.
J. Cell Biol.
99
,
2175
2186
7
de Forges
,
H.
,
Bouissou
,
A.
and
Perez
,
F.
(
2012
)
Interplay between microtubule dynamics and intracellular organization
.
Int. J. Biochem. Cell Biol.
44
,
266
274
8
Belmont
,
L.D.
,
Hyman
,
A.A.
,
Sawin
,
K.E.
and
Mitchison
,
T.J.
(
1990
)
Real-time visualization of cell cycle-dependent changes in microtubule dynamics in cytoplasmic extracts
.
Cell
62
,
579
589
9
Tamir
,
A.
,
Elad
,
N.
and
Medalia
,
O.
(
2011
)
Assembly and breakdown of microtubules within the midbody
.
Commun. Integr. Biol.
4
,
552
553
10
Raff
,
E.C.
,
Hoyle
,
H.D.
,
Popodi
,
E.M.
and
Turner
,
F.R.
(
2008
)
Axoneme β-tubulin sequence determines attachment of outer dynein arms
.
Curr. Biol.
18
,
911
914
11
Carvalho-Santos
,
Z.
,
Azimzadeh
,
J.
,
Pereira-Leal
,
J.B.
and
Bettencourt-Dias
,
M.
(
2011
)
Tracing the origins of centrioles, cilia, and flagella
.
J. Cell Biol.
194
,
165
175
12
Muroyama
,
A.
and
Lechler
,
T.
(
2017
)
Microtubule organization, dynamics and functions in differentiated cells
.
Development
144
,
3012
3021
13
Schwer
,
H.D.
,
Lecine
,
P.
,
Tiwari
,
S.
,
Italiano
,
J.E.J.
,
Hartwig
,
J.H.
and
Shivdasani
,
R.A.
(
2001
)
A lineage-restricted and divergent β-tubulin isoform is essential for the biogenesis, structure and function of blood platelets
.
Curr. Biol.
11
,
579
586
14
Patel-Hett
,
S.
,
Richardson
,
J.L.
,
Schulze
,
H.
,
Drabek
,
K.
,
Isaac
,
N.A.
,
Hoffmeister
,
K.
, et al.  (
2008
)
Visualization of microtubule growth in living platelets reveals a dynamic marginal band with multiple microtubules
.
Blood
111
,
4605
4616
15
Park
,
J.H.
and
Roll-Mecak
,
A.
(
2018
)
The tubulin code in neuronal polarity
.
Curr. Opin. Neurobiol.
51
,
95
102
16
Schulze
,
E.
and
Kirschner
,
M.
(
1987
)
Dynamic and stable populations of microtubules in cells
.
J. Cell Biol.
104
,
277
288
17
Kavallaris
,
M.
,
Don
,
S.
and
Verrills
,
N.
(
2008
) Microtubule-associated proteins and microtubule-interacting proteins: regulators of microtubule dynamics. In
The Role of Microtubules in Cell Biology, Neurobiology, and Oncology
(
Fojo
,
T.
, ed.), pp.
83
104
,
The Humana Press
,
Totowa, NJ
18
Ludueña
,
R.F.
(
2013
)
A hypothesis on the origin and evolution of tubulin
.
Int. Rev. Cell Mol. Biol.
302
,
41
185
19
Nogales
,
E.
,
Wolf
,
S.G.
and
Downing
,
K.H.
(
1998
)
Structure of the αβ tubulin dimer by electron crystallography
.
Nature
391
,
199
203
20
Ludueña
,
R.F.
(
1998
)
Multiple forms of tubulin: different gene products and covalent modifications
.
Int. Rev. Cytol.
178
,
207
275
21
Ludueña
,
R.F.
(
1993
)
Are tubulin isotypes functionally significant
.
Mol. Biol. Cell
4
,
445
457
22
Behnke
,
O.
and
Forer
,
A.
(
1967
)
Evidence for four classes of microtubules in individual cells
.
J. Cell Sci.
2
,
169
192
PMID:
[PubMed]
23
Fulton
,
C.
and
Simpson
,
P. A
. (
1976
) Selective synthesis and utilization of flagellar tubulin. The multi-tubulin hypothesis. In
Cell Motility: Cold Spring Harbor Conferences on Cell Proliferation
(
Goldman
,
R.
,
Pollard
,
T.
, and
Rosenbaum
,
J.
, eds.), pp.
987
1005
,
Cold Spring Harbor Press
,
New York
24
Cleveland
,
D.W.
,
Kirschner
,
M.W.
and
Cowan
,
N.J.
(
1978
)
Isolation of separate mRNAs for α- and β-tubulin and characterization of the corresponding in vitro translation products
.
Cell
15
,
1021
1031
25
Roll-Mecak
,
A.
(
2015
)
Intrinsically disordered tubulin tails: complex tuners of microtubule functions?
Semin. Cell Dev. Biol.
37
,
11
19
26
Jensen-Smith
,
H.C.
,
Ludueña
,
R.F.
and
Hallworth
,
R.
(
2003
)
Requirement for the βI and βIV tubulin isotypes in mammalian cilia
.
Cell Motil. Cytoskeleton
55
,
213
220
27
Guo
,
J.
,
Walss-Bass
,
C.
and
Ludueña
,
R.F.
(
2010
)
The β isotypes of tubulin in neuronal differentiation
.
Cytoskeleton
67
,
431
441
28
Hersheson
,
J.
,
Mencacci
,
N.E.
,
Davis
,
M.
,
MacDonald
,
N.
,
Trabzuni
,
D.
,
Ryten
,
M.
, et al.  (
2013
)
Mutations in the autoregulatory domain of β-tubulin 4a cause hereditary dystonia
.
Ann. Neurol.
73
,
546
553
29
Leandro-García
,
L.J.
,
Leskelä
,
S.
,
Landa
,
I.
,
Montero-Conde
,
C.
,
López-Jiménez
,
E.
,
Letón
,
R.
et al.  (
2010
)
Tumoral and tissue-specific expression of the major human β-tubulin isotypes
.
Cytoskeleton
67
,
214
223
30
Poirier
,
K.
,
Keays
,
D.A.
,
Francis
,
F.
,
Saillour
,
Y.
,
Bahi
,
N.
,
Manouvrier
,
S.
, et al.  (
2007
)
Large spectrum of lissencephaly and pachygyria phenotypes resulting from de novo missense mutations in tubulin alpha 1A (TUBA1A)
.
Hum. Mutat.
28
,
1055
1064
31
Petrukhin
,
K.E.
,
Speer
,
M.C.
,
Cayanis
,
E.
,
Bonaldo
,
M.F.
,
Tantravahi
,
U.
,
Soares
,
M.B.
et al.  (
1993
)
A microsatellite genetic linkage map of human chromosome 13
.
Genomics
15
,
76
85
32
Uhlen
,
M.
,
Fagerberg
,
L.
,
Hallstrom
,
B.M.
,
Lindskog
,
C.
,
Oksvold
,
P.
,
Mardinoglu
,
A.
, et al.  (
2015
)
Proteomics. Tissue-based map of the human proteome
.
Science
347
,
1260419
33
Thul
,
P.J.
,
Akesson
,
L.
,
Wiking
,
M.
,
Mahdessian
,
D.
,
Geladaki
,
A.
,
Ait Blal
,
H.
, et al.  (
2017
)
A subcellular map of the human proteome
.
Science
356
,
eaal13321
34
Stanchi
,
F.
,
Corso
,
V.
,
Scannapieco
,
P.
,
Ievolella
,
C.
,
Negrisolo
,
E.
,
Tiso
,
N.
et al.  (
2000
)
TUBA8: a new tissue-specific isoform of α-tubulin that is highly conserved in human and mouse
.
Biochem. Biophys. Res. Commun.
270
,
1111
1118
35
Braun
,
A.
,
Breuss
,
M.
,
Salzer
,
M.C.
,
Flint
,
J.
,
Cowan
,
N.J.
and
Keays
,
D.A.
(
2010
)
Tuba8 is expressed at low levels in the developing mouse and human brain
.
Am. J. Hum. Genet.
86
,
819
822
36
Feng
,
R.
,
Sang
,
Q.
,
Kuang
,
Y.
,
Sun
,
X.
,
Yan
,
Z.
,
Zhang
,
S.
, et al.  (
2016
)
Mutations in TUBB8 and human oocyte meiotic arrest
.
N. Engl. J. Med.
374
,
223
232
37
Katsetos
,
C.D.
,
Legido
,
A.
,
Perentes
,
E.
and
Mörk
,
S.J.
(
2003
)
Class III β-tubulin isotype: a key cytoskeletal protein at the crossroads of developmental neurobiology and tumor neuropathology
.
J. Child Neurol.
18
,
851
866
;
discussion 867
38
Lewis
,
S.A.
,
Gu
,
W.
and
Cowan
,
N.J.
(
1987
)
Free intermingling of mammalian β-tubulin isotypes among functionally distinct microtubules
.
Cell
49
,
539
548
39
Hoyle
,
H.D.
and
Raff
,
E.C.
(
1990
)
Two Drosophila beta tubulin isoforms are not functionally equivalent
.
J. Cell Biol.
111
,
1009
1026
40
Saillour
,
Y.
,
Broix
,
L.
,
Bruel-Jungerman
,
E.
,
Lebrun
,
N.
,
Muraca
,
G.
,
Rucci
,
J.
et al.  (
2014
)
Beta tubulin isoforms are not interchangeable for rescuing impaired radial migration due to Tubb3 knockdown
.
Hum. Mol. Genet.
23
,
1516
1526
41
Song
,
Y.
and
Brady
,
S.T.
(
2015
)
Post-translational modifications of tubulin: pathways to functional diversity of microtubules
.
Trends Cell Biol.
25
,
125
136
42
Janke
,
C.
(
2014
)
The tubulin code: molecular components, readout mechanisms, and functions
.
J. Cell Biol.
206
,
461
472
43
Wloga
,
D.
,
Joachimiak
,
E.
and
Fabczak
,
H.
(
2017
)
Tubulin post-translational modifications and microtubule dynamics
.
Int. J. Mol. Sci.
18
,
e2207
44
Yu
,
I.
,
Garnham
,
C.P.
and
Roll-Mecak
,
A.
(
2015
)
Writing and reading the tubulin code
.
J. Biol. Chem.
290
,
17163
17172
45
Soppina
,
V.
,
Herbstman
,
J.F.
,
Skiniotis
,
G.
and
Verhey
,
K.J.
(
2012
)
Luminal localization of α-tubulin K40 acetylation by cryo-EM analysis of Fab-labeled microtubules
.
PLoS ONE
7
,
e48204
46
Fourest-Lieuvin
,
A.
,
Peris
,
L.
,
Gache
,
V.
,
Garcia-Saez
,
I.
,
Juillan-Binard
,
C.
,
Lantez
,
V.
et al.  (
2006
)
Microtubule regulation in mitosis: tubulin phosphorylation by the cyclin-dependent kinase Cdk1
.
Mol. Biol. Cell
17
,
1041
1050
47
Song
,
Y.
,
Kirkpatrick
,
L.L.
,
Schilling
,
A.B.
,
Helseth
,
D.L.
,
Chabot
,
N.
,
Keillor
,
J.W.
et al.  (
2013
)
Transglutaminase and polyamination of tubulin: posttranslational modification for stabilizing axonal microtubules
.
Neuron
78
,
109
123
48
Park
,
I.Y.
,
Chowdhury
,
P.
,
Tripathi
,
D.N.
,
Powell
,
R.T.
,
Dere
,
R.
,
Terzo
,
E.A.
et al.  (
2016
)
Methylated α-tubulin antibodies recognize a new microtubule modification on mitotic microtubules
.
mAbs
8
,
1590
1597
49
Coombes
,
C.
,
Yamamoto
,
A.
,
McClellan
,
M.
,
Reid
,
T.A.
,
Plooster
,
M.
,
Luxton
,
G.W.G.
et al.  (
2016
)
Mechanism of microtubule lumen entry for the α-tubulin acetyltransferase enzyme αTAT1
.
Proc. Natl Acad. Sci. U.S.A.
113
,
E7176
E7184
50
Xu
,
Z.
,
Schaedel
,
L.
,
Portran
,
D.
,
Aguilar
,
A.
,
Gaillard
,
J.
,
Marinkovich
,
M.P.
et al.  (
2017
)
Microtubules acquire resistance from mechanical breakage through intralumenal acetylation
.
Science
356
,
328
332
51
Peris
,
L.
,
Thery
,
M.
,
Fauré
,
J.
,
Saoudi
,
Y.
,
Lafanechère
,
L.
,
Chilton
,
J.K.
, et al.  (
2006
)
Tubulin tyrosination is a major factor affecting the recruitment of CAP-Gly proteins at microtubule plus ends
.
J. Cell Biol.
174
,
839
849
52
Barisic
,
M.
,
Silva e Sousa
,
R.
,
Tripathy
,
S.K.
,
Magiera
,
M.M.
,
Zaytsev
,
A.V.
,
Pereira
,
A.L.
et al.  (
2015
)
Mitosis. Microtubule detyrosination guides chromosomes during mitosis
.
Science
348
,
799
803
53
Tas
,
R.P.
,
Chazeau
,
A.
,
Cloin
,
B.M.C.
,
Lambers
,
M.L.A.
,
Hoogenraad
,
C.C.
and
Kapitein
,
L.C.
(
2017
)
Differentiation between oppositely oriented microtubules controls polarized neuronal transport
.
Neuron
96
,
1264
1271.e5
54
Verhey
,
K.J.
and
Gaertig
,
J.
(
2007
)
The tubulin code
.
Cell Cycle
6
,
2152
2160
55
Janke
,
C.
and
Bulinski
,
J.C.
(
2011
)
Post-translational regulation of the microtubule cytoskeleton: mechanisms and functions
.
Nat. Rev. Mol. Cell Biol.
12
,
773
786
56
Panda
,
D.
,
Miller
,
H.P.
,
Banerjee
,
A.
,
Ludueña
,
R.F.
and
Wilson
,
L.
(
1994
)
Microtubule dynamics in vitro are regulated by the tubulin isotype composition
.
Proc. Natl Acad. Sci. U.S.A.
91
,
11358
11362
57
Vemu
,
A.
,
Atherton
,
J.
,
Spector
,
J.O.
,
Moores
,
C.A.
and
Roll-Mecak
,
A.
(
2017
)
Tubulin isoform composition tunes microtubule dynamics
.
Mol. Biol. Cell
28
,
3564
3572
58
Ti
,
S.-C.
,
Alushin
,
G.M.
and
Kapoor
,
T.M.
(
2018
)
Human β-tubulin isotypes can regulate microtubule protofilament number and stability
.
Dev. Cell
47
,
175
190.e5
59
Nielsen
,
M.G.
,
Turner
,
F.R.
,
Hutchens
,
J.A.
and
Raff
,
E.C.
(
2001
)
Axoneme-specific β-tubulin specialization: a conserved C-terminal motif specifies the central pair
.
Curr. Biol.
11
,
529
533
60
Wang
,
D.
,
Villasante
,
A.
,
Lewis
,
S.A.
and
Cowan
,
N.J.
(
1986
)
The mammalian β-tubulin repertoire: hematopoietic expression of a novel, heterologous β-tubulin isotype
.
J. Cell Biol.
103
,
1903
1910
61
Magiera
,
M.M.
,
Singh
,
P.
and
Janke
,
C.
(
2018
)
Snapshot: functions of tubulin posttranslational modifications
.
Cell
173
,
1552
1552.e1
62
Portran
,
D.
,
Schaedel
,
L.
,
Xu
,
Z.
,
Théry
,
M.
and
Nachury
,
M.V.
(
2017
)
Tubulin acetylation protects long-lived microtubules against mechanical aging
.
Nat. Cell Biol.
19
,
391
398
63
Valenstein
,
M.L.
and
Roll-Mecak
,
A.
(
2016
)
Graded control of microtubule severing by tubulin glutamylation
.
Cell
164
,
911
921
64
Kubo
,
T.
,
Yanagisawa
,
H.
,
Yagi
,
T.
,
Hirono
,
M.
and
Kamiya
,
R.
(
2010
)
Tubulin polyglutamylation regulates axonemal motility by modulating activities of inner-arm dyneins
.
Curr. Biol.
20
,
441
445
65
Suryavanshi
,
S.
,
Eddé
,
B.
,
Fox
,
L.A.
,
Guerrero
,
S.
,
Hard
,
R.
,
Hennessey
,
T.
, et al.  (
2010
)
Tubulin glutamylation regulates ciliary motility by altering inner dynein arm activity
.
Curr. Biol.
20
,
435
440
66
Gadadhar
,
S.
,
Dadi
,
H.
,
Bodakuntla
,
S.
,
Schnitzler
,
A.
,
Bieche
,
I.
,
Rusconi
,
F.
et al.  (
2017
)
Tubulin glycylation controls primary cilia length
.
J. Cell Biol.
216
,
2701
2713
67
Peris
,
L.
,
Wagenbach
,
M.
,
Lafanechère
,
L.
,
Brocard
,
J.
,
Moore
,
A.T.
,
Kozielski
,
F.
et al.  (
2009
)
Motor-dependent microtubule disassembly driven by tubulin tyrosination
.
J. Cell Biol.
185
,
1159
1166
68
Konno
,
A.
,
Setou
,
M.
and
Ikegami
,
K.
(
2012
)
Ciliary and flagellar structure and function – their regulations by posttranslational modifications of axonemal tubulin
.
Int. Rev. Cell Mol. Biol.
294
,
133
170
69
Konishi
,
Y.
and
Setou
,
M.
(
2009
)
Tubulin tyrosination navigates the kinesin-1 motor domain to axons
.
Nat. Neurosci.
12
,
559
70
Liao
,
G.
and
Gundersen
,
G.G.
(
1998
)
Kinesin is a candidate for cross-bridging microtubules and intermediate filaments. Selective binding of kinesin to detyrosinated tubulin and vimentin
.
J. Biol. Chem.
273
,
9797
9803
71
Dunn
,
S.
,
Morrison
,
E.E.
,
Liverpool
,
T.B.
,
Molina-París
,
C.
,
Cross
,
R.A.
,
Alonso
,
M.C.
et al.  (
2008
)
Differential trafficking of Kif5c on tyrosinated and detyrosinated microtubules in live cells
.
J. Cell Sci.
121
,
1085
1095
72
Sirajuddin
,
M.
,
Rice
,
L.M.
and
Vale
,
R.D.
(
2014
)
Regulation of microtubule motors by tubulin isotypes and post-translational modifications
.
Nat. Cell Biol.
16
,
335
344
73
Reed
,
N.A.
,
Cai
,
D.
,
Blasius
,
T.L.
,
Jih
,
G.T.
,
Meyhofer
,
E.
,
Gaertig
,
J.
et al.  (
2006
)
Microtubule acetylation promotes kinesin-1 binding and transport
.
Curr. Biol.
16
,
2166
2172
74
McKenney
,
R.J.
,
Huynh
,
W.
,
Vale
,
R.D.
and
Sirajuddin
,
M.
(
2016
)
Tyrosination of α-tubulin controls the initiation of processive dynein-dynactin motility
.
EMBO J.
35
,
1175
1185
75
Muñoz-Castañeda
,
R.
,
Díaz
,
D.
,
Peris
,
L.
,
Andrieux
,
A.
,
Bosc
,
C.
,
Muñoz-Castañeda
,
J.M.
et al.  (
2018
)
Cytoskeleton stability is essential for the integrity of the cerebellum and its motor- and affective-related behaviors
.
Sci. Rep.
8
,
3072
76
Erck
,
C.
,
Peris
,
L.
,
Andrieux
,
A.
,
Meissirel
,
C.
,
Gruber
,
A.D.
,
Vernet
,
M.
, et al.  (
2005
)
A vital role of tubulin-tyrosine-ligase for neuronal organization
.
Proc. Natl Acad. Sci. U.S.A.
102
,
7853
7858
77
Marcos
,
S.
,
Moreau
,
J.
,
Backer
,
S.
,
Job
,
D.
,
Andrieux
,
A.
and
Bloch-Gallego
,
E.
(
2009
)
Tubulin tyrosination is required for the proper organization and pathfinding of the growth cone
.
PLoS ONE
4
,
e5405
78
Jordan
,
M.A.
and
Wilson
,
L.
(
2004
)
Microtubules as a target for anticancer drugs
.
Nat. Rev. Cancer
4
,
253
265
79
Dumontet
,
C.
and
Jordan
,
M.A.
(
2010
)
Microtubule-binding agents: a dynamic field of cancer therapeutics
.
Nat. Rev. Drug Discov.
9
,
790
803
80
Cao
,
Y.-N.
,
Zheng
,
L.-L.
,
Wang
,
D.
,
Liang
,
X.-X.
,
Gao
,
F.
and
Zhou
,
X.-L.
(
2018
)
Recent advances in microtubule-stabilizing agents
.
Eur. J. Med. Chem.
143
,
806
828
81
Lu
,
C.
,
Zhang
,
J.
,
He
,
S.
,
Wan
,
C.
,
Shan
,
A.
,
Wang
,
Y.
, et al.  (
2013
)
Increased α-tubulin1b expression indicates poor prognosis and resistance to chemotherapy in hepatocellular carcinoma
.
Dig. Dis. Sci.
58
,
2713
2720
82
Hasegawa
,
S.
,
Miyoshi
,
Y.
,
Egawa
,
C.
,
Ishitobi
,
M.
,
Taguchi
,
T.
,
Tamaki
,
Y.
et al.  (
2003
)
Prediction of response to docetaxel by quantitative analysis of class I and III β-tubulin isotype mRNA expression in human breast cancers
.
Clin. Cancer Res.
9
,
2992
2997
PMID:
[PubMed]
83
Kavallaris
,
M.
,
Kuo
,
D.Y.
,
Burkhart
,
C.A.
,
Regl
,
D.L.
,
Norris
,
M.D.
,
Haber
,
M.
et al.  (
1997
)
Taxol-resistant epithelial ovarian tumors are associated with altered expression of specific beta-tubulin isotypes
.
J. Clin. Invest.
100
,
1282
1293
84
Cucchiarelli
,
V.
,
Hiser
,
L.
,
Smith
,
H.
,
Frankfurter
,
A.
,
Spano
,
A.
,
Correia
,
J.J.
et al.  (
2008
)
β-Tubulin isotype classes II and V expression patterns in nonsmall cell lung carcinomas
.
Cell Motil. Cytoskeleton
65
,
675
685
85
Christoph
,
D.C.
,
Kasper
,
S.
,
Gauler
,
T.C.
,
Loesch
,
C.
,
Engelhard
,
M.
,
Theegarten
,
D.
, et al.  (
2012
)
βV-Tubulin expression is associated with outcome following taxane-based chemotherapy in non-small cell lung cancer
.
Br. J. Cancer
107
,
823
830
86
Blenk
,
S.
,
Engelmann
,
J.C.
,
Pinkert
,
S.
,
Weniger
,
M.
,
Schultz
,
J.
,
Rosenwald
,
A.
et al.  (
2008
)
Explorative data analysis of MCL reveals gene expression networks implicated in survival and prognosis supported by explorative CGH analysis
.
BMC Cancer
8
,
106
87
Cullen
,
K.J.
,
Schumaker
,
L.
,
Nikitakis
,
N.
,
Goloubeva
,
O.
,
Tan
,
M.
,
Sarlis
,
N.J.
et al.  (
2009
)
β-Tubulin-II expression strongly predicts outcome in patients receiving induction chemotherapy for locally advanced squamous carcinoma of the head and neck: a companion analysis of the TAX 324 trial
.
J. Clin. Oncol.
27
,
6222
6228
88
Paradiso
,
A.
,
Mangia
,
A.
,
Chiriatti
,
A.
,
Tommasi
,
S.
,
Zito
,
A.
,
Latorre
,
A.
et al.  (
2005
)
Biomarkers predictive for clinical efficacy of taxol-based chemotherapy in advanced breast cancer
.
Ann. Oncol.
16
,
iv14
iv19
89
Gan
,
P.P.
,
McCarroll
,
J.A.
,
Byrne
,
F.L.
,
Garner
,
J.
and
Kavallaris
,
M.
(
2011
)
Specific β-tubulin isotypes can functionally enhance or diminish epothilone B sensitivity in non-small cell lung cancer cells
.
PLoS ONE
6
,
e21717
90
Hwang
,
J.-E.
,
Hong
,
J.-Y.
,
Kim
,
K.
,
Kim
,
S.-H.
,
Choi
,
W.-Y.
,
Kim
,
M.-J.
, et al.  (
2013
)
Class III β-tubulin is a predictive marker for taxane-based chemotherapy in recurrent and metastatic gastric cancer
.
BMC Cancer
13
,
431
91
Ploussard
,
G.
,
Terry
,
S.
,
Maille
,
P.
,
Allory
,
Y.
,
Sirab
,
N.
,
Kheuang
,
L.
, et al.  (
2010
)
Class III β-tubulin expression predicts prostate tumor aggressiveness and patient response to docetaxel-based chemotherapy
.
Cancer Res.
70
,
9253
9264
92
Roque
,
D.M.
,
Bellone
,
S.
,
English
,
D.P.
,
Buza
,
N.
,
Cocco
,
E.
,
Gasparrini
,
S.
, et al.  (
2013
)
Tubulin-β-III overexpression by uterine serous carcinomas is a marker for poor overall survival after platinum/taxane chemotherapy and sensitivity to epothilones
.
Cancer
119
,
2582
2592
93
Nienstedt
,
J.C.
,
Grobe
,
A.
,
Clauditz
,
T.
,
Simon
,
R.
,
Muenscher
,
A.
,
Knecht
,
R.
, et al.  (
2017
)
High-level βIII-tubulin overexpression occurs in most head and neck cancers but is unrelated to clinical outcome
.
J. Oral Pathol. Med.
46
,
986
990
94
Hasegawa
,
S.
,
Miyoshi
,
Y.
,
Egawa
,
C.
,
Ishitobi
,
M.
,
Tamaki
,
Y.
,
Monden
,
M.
et al.  (
2002
)
Mutational analysis of the class I β-tubulin gene in human breast cancer
.
Int. J. Cancer
101
,
46
51
95
Ohishi
,
Y.
,
Oda
,
Y.
,
Basaki
,
Y.
,
Kobayashi
,
H.
,
Wake
,
N.
,
Kuwano
,
M.
et al.  (
2007
)
Expression of β-tubulin isotypes in human primary ovarian carcinoma
.
Gynecol. Oncol.
105
,
586
592
96
Bernard-Marty
,
C.
,
Treilleux
,
I.
,
Dumontet
,
C.
,
Cardoso
,
F.
,
Fellous
,
A.
,
Gancberg
,
D.
, et al.  (
2002
)
Microtubule-associated parameters as predictive markers of docetaxel activity in advanced breast cancer patients: results of a pilot study
.
Clin. Breast Cancer
3
,
341
345
97
Shalli
,
K.
,
Brown
,
I.
,
Heys
,
S.D.
and
Schofield
,
A.C.
(
2005
)
Alterations of β-tubulin isotypes in breast cancer cells resistant to docetaxel
.
FASEB J.
19
,
1299
1301
98
Mathew
,
D.
,
Wang
,
Y.
,
Van Arsdale
,
A.
,
Horwitz
,
S.B.
and
McDaid
,
H.
(
2018
)
Expression of βV-tubulin in secretory cells of the fallopian tube epithelium marks cellular atypia
.
Int. J. Gynecol. Cancer
28
,
363
370
99
Chao
,
S.K.
,
Wang
,
Y.
,
Verdier-Pinard
,
P.
,
Yang
,
C.-P.H.
,
Liu
,
L.
,
Rodriguez-Gabin
,
A.
et al.  (
2012
)
Characterization of a human βV-tubulin antibody and expression of this isotype in normal and malignant human tissue
.
Cytoskeleton
69
,
566
576
100
Parker
,
A.L.
,
Teo
,
W.S.
,
McCarroll
,
J.A.
and
Kavallaris
,
M.
(
2017
)
An emerging role for tubulin isotypes in modulating cancer biology and chemotherapy resistance
.
Int. J. Mol. Sci.
18
,
E1434
101
Gan
,
P.P.
and
Kavallaris
,
M.
(
2008
)
Tubulin-targeted drug action: functional significance of class ii and class IVb β-tubulin in vinca alkaloid sensitivity
.
Cancer Res.
68
,
9817
9824
102
McCarroll
,
J.A.
,
Sharbeen
,
G.
,
Liu
,
J.
,
Youkhana
,
J.
,
Goldstein
,
D.
,
McCarthy
,
N.
, et al.  (
2015
)
βIII-tubulin: a novel mediator of chemoresistance and metastases in pancreatic cancer
.
Oncotarget
6
,
2235
2249
103
Derry
,
W.B.
,
Wilson
,
L.
,
Khan
,
I.A.
,
Ludueña
,
R.F.
and
Jordan
,
M.A.
(
1997
)
Taxol differentially modulates the dynamics of microtubules assembled from unfractionated and purified β-tubulin isotypes
.
Biochemistry
36
,
3554
3562
104
Gan
,
P.P.
,
Pasquier
,
E.
and
Kavallaris
,
M.
(
2007
)
Class III β-tubulin mediates sensitivity to chemotherapeutic drugs in non small cell lung cancer
.
Cancer Res.
67
,
9356
9363
105
Rai
,
A.
,
Kapoor
,
S.
,
Naaz
,
A.
,
Kumar Santra
,
M.
and
Panda
,
D.
(
2017
)
Enhanced stability of microtubules contributes in the development of colchicine resistance in MCF-7 cells
.
Biochem. Pharmacol.
132
,
38
47
106
Ranganathan
,
S.
,
McCauley
,
R.A.
,
Dexter
,
D.W.
and
Hudes
,
G.R.
(
2001
)
Modulation of endogenous β-tubulin isotype expression as a result of human β(III)cDNA transfection into prostate carcinoma cells
.
Br. J. Cancer
85
,
735
740
107
Aoki
,
D.
,
Oda
,
Y.
,
Hattori
,
S.
,
Taguchi
,
K.
,
Ohishi
,
Y.
,
Basaki
,
Y.
, et al.  (
2009
)
Overexpression of class III β-tubulin predicts good response to taxane-based chemotherapy in ovarian clear cell adenocarcinoma
.
Clin. Cancer Res.
15
,
1473
1480
108
Kanakkanthara
,
A.
,
Northcote
,
P.T.
and
Miller
,
J.H.
(
2012
)
βII-tubulin and βIII-tubulin mediate sensitivity to peloruside A and laulimalide, but not paclitaxel or vinblastine, in human ovarian carcinoma cells
.
Mol. Cancer Ther.
11
,
393
404
109
Lafanechère
,
L.
,
Courtay-Cahen
,
C.
,
Kawakami
,
T.
,
Jacrot
,
M.
,
Rudiger
,
M.
,
Wehland
,
J.
et al.  (
1998
)
Suppression of tubulin tyrosine ligase during tumor growth
.
J. Cell Sci.
111
,
171
181
PMID:
[PubMed]
110
Souček
,
K.
,
Kamaid
,
A.
,
Phung
,
A.D.
,
Kubala
,
L.
,
Bulinski
,
J.C.
,
Harper
,
R.W.
et al.  (
2006
)
Normal and prostate cancer cells display distinct molecular profiles of α-tubulin posttranslational modifications
.
Prostate
66
,
954
965
111
Mialhe
,
A.
,
Lafanechère
,
L.
,
Treilleux
,
I.
,
Peloux
,
N.
,
Dumontet
,
C.
,
Brémond
,
A.
, et al.  (
2001
)
Tubulin detyrosination is a frequent occurrence in breast cancers of poor prognosis
.
Cancer Res.
61
,
5024
5027
PMID:
[PubMed]
112
Kato
,
C.
,
Miyazaki
,
K.
,
Nakagawa
,
A.
,
Ohira
,
M.
,
Nakamura
,
Y.
,
Ozaki
,
T.
et al.  (
2004
)
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
,
365
375
113
Whipple
,
R.A.
,
Matrone
,
M.A.
,
Cho
,
E.H.
,
Balzer
,
E.M.
,
Vitolo
,
M.I.
,
Yoon
,
J.R.
et al.  (
2010
)
Epithelial-to-mesenchymal transition promotes tubulin detyrosination and microtentacles that enhance endothelial engagement
.
Cancer Res.
70
,
8127
8137
114
Zhang
,
B.
,
Wu
,
Z.
,
Xie
,
W.
,
Tian
,
D.
,
Chen
,
F.
,
Qin
,
C.
, et al.  (
2017
)
The expression of vasohibin-1 and its prognostic significance in bladder cancer
.
Exp. Ther. Med.
14
,
3477
3484
115
Aillaud
,
C.
,
Bosc
,
C.
,
Peris
,
L.
,
Bosson
,
A.
,
Heemeryck
,
P.
,
Van Dijk
,
J.
, et al.  (
2017
)
Vasohibins/SVBP are tubulin carboxypeptidases (TCPs) that regulate neuron differentiation
.
Science
358
,
1448
1453
116
Giustiniani
,
J.
,
Daire
,
V.
,
Cantaloube
,
I.
,
Durand
,
G.
,
Poüs
,
C.
,
Perdiz
,
D.
et al.  (
2009
)
Tubulin acetylation favors Hsp90 recruitment to microtubules and stimulates the signaling function of the Hsp90 clients Akt/PKB and p53
.
Cell. Signal.
21
,
529
539
117
Zhang
,
Z.
,
Yamashita
,
H.
,
Toyama
,
T.
,
Sugiura
,
H.
,
Omoto
,
Y.
,
Ando
,
Y.
et al.  (
2004
)
HDAC6 expression is correlated with better survival in breast cancer
.
Clin. Cancer Res.
10
,
6962
6968
118
Asthana
,
J.
,
Kapoor
,
S.
,
Mohan
,
R.
and
Panda
,
D.
(
2013
)
Inhibition of HDAC6 deacetylase activity increases its binding with microtubules and suppresses microtubule dynamic instability in MCF-7 cells
.
J. Biol. Chem.
288
,
22516
22526
119
Sangrajrang
,
S.
and
Fellous
,
A.
(
2000
)
Taxol resistance
.
Chemotherapy
46
,
327
334
120
Rocha
,
C.
,
Papon
,
L.
,
Cacheux
,
W.
,
Marques Sousa
,
P.
,
Lascano
,
V.
,
Tort
,
O.
, et al.  (
2014
)
Tubulin glycylases are required for primary cilia, control of cell proliferation and tumor development in colon
.
EMBO J.
33
,
2247
2260
121
Miller
,
L.M.
,
Menthena
,
A.
,
Chatterjee
,
C.
,
Verdier-Pinard
,
P.
,
Novikoff
,
P.M.
,
Horwitz
,
S.B.
et al.  (
2008
)
Increased levels of a unique post-translationally modified βIVb-tubulin isotype in liver cancer
.
Biochemistry
47
,
7572
7582
122
Kavallaris
,
M.
(
2010
)
Microtubules and resistance to tubulin-binding agents
.
Nat. Rev. Cancer
10
,
194
204
123
Zhou
,
J.
,
Gupta
,
K.
,
Yao
,
J.
,
Ye
,
K.
,
Panda
,
D.
,
Giannakakou
,
P.
et al.  (
2002
)
Paclitaxel-resistant human ovarian cancer cells undergo c-Jun NH2-terminal kinase-mediated apoptosis in response to noscapine
.
J. Biol. Chem.
277
,
39777
39785
124
Pryor
,
D.E.
,
O'Brate
,
A.
,
Bilcer
,
G.
,
Diaz
,
J.F.
,
Wang
,
Y.
,
Wang
,
Y.
, et al.  (
2002
)
The microtubule stabilizing agent laulimalide does not bind in the taxoid site, kills cells resistant to paclitaxel and epothilones, and may not require its epoxide moiety for activity
.
Biochemistry
41
,
9109
9115
125
Gaitanos
,
T.N.
,
Buey
,
R.M.
,
Diaz
,
J.F.
,
Northcote
,
P.T.
,
Teesdale-Spittle
,
P.
,
Andreu
,
J.M.
et al.  (
2004
)
Peloruside A does not bind to the taxoid site on β-tubulin and retains its activity in multidrug-resistant cell lines
.
Cancer Res.
64
,
5063
5067
126
Risinger
,
A.L.
,
Jackson
,
E.M.
,
Polin
,
L.A.
,
Helms
,
G.L.
,
LeBoeuf
,
D.A.
,
Joe
,
P.A.
et al.  (
2008
)
The taccalonolides: microtubule stabilizers that circumvent clinically relevant taxane resistance mechanisms
.
Cancer Res.
68
,
8881
8888
127
Field
,
J.J.
,
Northcote
,
P.T.
,
Paterson
,
I.
,
Altmann
,
K.-H.
,
Díaz
,
J.F.
and
Miller
,
J.H.
(
2017
)
Zampanolide, a microtubule-stabilizing agent, is active in resistant cancer cells and inhibits cell migration
.
Int. J. Mol. Sci.
18
,
E971
128
Matesanz
,
R.
,
Trigili
,
C.
,
Rodriguez-Salarichs
,
J.
,
Zanardi
,
I.
,
Pera
,
B.
,
Nogales
,
A.
, et al.  (
2014
)
Taxanes with high potency inducing tubulin assembly overcome tumoural cell resistances
.
Bioorg. Med. Chem.
22
,
5078
5090
129
Ferlini
,
C.
,
Raspaglio
,
G.
,
Mozzetti
,
S.
,
Cicchillitti
,
L.
,
Filippetti
,
F.
,
Gallo
,
D.
et al.  (
2005
)
The seco-taxane IDN5390 is able to target class III β-tubulin and to overcome paclitaxel resistance
.
Cancer Res.
65
,
2397
2405
130
Dumontet
,
C.
,
Jordan
,
M.A.
and
Lee
,
F.F.Y.
(
2009
)
Ixabepilone: targeting βIII-tubulin expression in taxane-resistant malignancies
.
Mol. Cancer Ther.
8
,
17
25
131
Smiyun
,
G.
,
Azarenko
,
O.
,
Miller
,
H.
,
Rifkind
,
A.
,
LaPointe
,
N.E.
,
Wilson
,
L.
et al.  (
2017
)
βIII-tubulin enhances efficacy of cabazitaxel as compared with docetaxel
.
Cancer Chemother. Pharmacol.
80
,
151
164
132
Yang
,
C.-P.H.
,
Yap
,
E.-H.
,
Xiao
,
H.
,
Fiser
,
A.
and
Horwitz
,
S.B.
(
2016
)
2-(m-Azidobenzoyl)taxol binds differentially to distinct β-tubulin isotypes
.
Proc. Natl Acad. Sci. U.S.A.
113
,
11294
11299
133
Radakovic
,
A.
and
Boger
,
D.L.
(
2018
)
High expression of class III β-tubulin has no impact on functional cancer cell growth inhibition of a series of key vinblastine analogs
.
Bioorg. Med. Chem. Lett.
28
,
863
865
134
Arnst
,
K.E.
,
Wang
,
Y.
,
Hwang
,
D.-J.
,
Xue
,
Y.
,
Costello
,
T.
,
Hamilton
,
D.
, et al.  (
2018
)
A potent, metabolically stable tubulin inhibitor targets the colchicine binding site and overcomes taxane resistance
.
Cancer Res.
78
,
265
277
135
Lindamulage
,
I.K.
,
Vu
,
H.-Y.
,
Karthikeyan
,
C.
,
Knockleby
,
J.
,
Lee
,
Y.-F.
,
Trivedi
,
P.
et al.  (
2017
)
Novel quinolone chalcones targeting colchicine-binding pocket kill multidrug-resistant cancer cells by inhibiting tubulin activity and MRP1 function
.
Sci. Rep.
7
,
10298
136
Denoulet
,
P.
,
Eddé
,
B.
and
Gros
,
F.
(
1986
)
Differential expression of several neurospecific β-tubulin mRNAs in the mouse brain during development
.
Gene
50
,
289
297
137
Paturle
,
L.
,
Wehland
,
J.
,
Margolis
,
R.L.
and
Job
,
D.
(
1989
)
Complete separation of tyrosinated, detyrosinated, and nontyrosinatable brain tubulin subpopulations using affinity chromatography
.
Biochemistry
28
,
2698
2704
138
Yaffe
,
M.B.
,
Levison
,
B.S.
,
Szasz
,
J.
and
Sternlicht
,
H.
(
1988
)
Expression of a human α-tubulin: properties of the isolated subunit
.
Biochemistry
27
,
1869
1880
139
Gao
,
Y.
,
Vainberg
,
I.E.
Chow
,
R.L.
and
Cowan
,
N.J.
(
1993
)
Two cofactors and cytoplasmic chaperonin are required for the folding of α- and β-tubulin
.
Mol. Cell. Biol.
13
,
2478
2485
140
Tian
,
G.
,
Huang
,
Y.
,
Rommelaere
,
H.
,
Vandekerckhove
,
J.
,
Ampe
,
C.
and
Cowan
,
N.J.
(
1996
)
Pathway leading to correctly folded β-tubulin
.
Cell
86
,
287
296
141
Weatherbee
,
J.A.
,
Luftig
,
R.B.
and
Weihing
,
R.R.
(
1980
)
Purification and reconstitution of HeLa cell microtubules
.
Biochemistry
19
,
4116
4123
142
Fourest-Lieuvin
,
A.
(
2006
)
Purification of tubulin from limited volumes of cultured cells
.
Protein Expr. Purif.
45
,
183
190
143
Vemu
,
A.
,
Garnham
,
C.P.
,
Lee
,
D.-Y.
and
Roll-Mecak
,
A.
(
2014
)
Generation of differentially modified microtubules using in vitro enzymatic approaches
.
Methods Enzymol.
540
,
149
166
144
Lacroix
,
B.
and
Janke
,
C.
(
2011
)
Generation of differentially polyglutamylated microtubules
.
Methods Mol. Biol.
777
,
57
69
145
Lacroix
,
B.
,
van Dijk
,
J.
,
Gold
,
N.D.
,
Guizetti
,
J.
,
Aldrian-Herrada
,
G.
,
Rogowski
,
K.
et al.  (
2010
)
Tubulin polyglutamylation stimulates spastin-mediated microtubule severing
.
J. Cell Biol.
189
,
945
954
146
Widlund
,
P.O.
,
Podolski
,
M.
,
Reber
,
S.
,
Alper
,
J.
,
Storch
,
M.
,
Hyman
,
A.A.
et al.  (
2012
)
One-step purification of assembly-competent tubulin from diverse eukaryotic sources
.
Mol. Biol. Cell
23
,
4393
4401
147
Alper
,
J.D.
,
Decker
,
F.
,
Agana
,
B.
and
Howard
,
J.
(
2014
)
The motility of axonemal dynein is regulated by the tubulin code
.
Biophys. J.
107
,
2872
2880
148
Drummond
,
D.R.
,
Kain
,
S.
,
Newcombe
,
A.
,
Hoey
,
C.
,
Katsuki
,
M.
and
Cross
,
R.A.
(
2011
)
Purification of tubulin from the fission yeast Schizosaccharomyces pombe
.
Methods Mol. Biol.
777
,
29
55
149
Vemu
,
A.
,
Atherton
,
J.
,
Spector
,
J.O.
,
Szyk
,
A.
,
Moores
,
C.A.
and
Roll-Mecak
,
A.
(
2016
)
Structure and dynamics of single-isoform recombinant neuronal human tubulin
.
J. Biol. Chem.
291
,
12907
12915