Self-organization is a recurrent theme in biology, evident across multiple length scales ranging from the oscillations of nanometer-sized signaling molecules in the cytoplasm to the formation of micron-sized cellular organelles and the patterning of millimeter-sized tissues. One of the best cellular systems for elucidating the principles of self-organization is the microtubule cytoskeleton, where dynamic microtubule polymers are organized into micron-sized arrays (Karsenti, 2008). Every microtubule filament within a multi-microtubule array is a polymer of tubulin subunits, polymerized with a specific polarity. Despite being assembled from the same microtubule building block, microtubule-based arrays display considerable diversity in their overall dimensions, shape, and polarity, in a manner that is intricately linked to their function. For instance, the orientation of coaligned, overlapping microtubules in the cortex of plant cells specifies the axis of cell elongation, and the polarity of long microtubule bundles in neurons distinguishes axons from dendrites (Chen et al., 2016; Kelliher et al., 2019). Understanding the mechanisms that direct the formation of distinct cellular arrays and specify their architecture will bring to light the general design principles underlying self-organization.

Building a multi-microtubule array of defined architecture requires the regulation of dimensions of individual polymers, as well as organizing them into higher-order structures. Microtubules are characterized by their inherent dynamic instability and cycle between phases of polymerization and depolymerization (Mitchison and Kirschner, 1984). The parameters governing dynamic instability therefore specify the stability, length, and density of microtubules within an array. There are broadly two ways in which dynamic instability is thought to be regulated. First, polymerization dynamics of individual microtubule filaments are sensitive to the concentration of soluble tubulin heterodimers which constitute their building blocks (Walker et al., 1988). In cells, tubulin levels are in turn autoregulated through translational control by tubulin itself (Cleveland, 1988; Lin et al., 2020). Second, the dynamics of microtubule polymers are regulated by the activity of microtubule-associated proteins (MAPs), which include both motor and non-motor proteins. In addition to polymer length and stability, MAPs also mediate the organization of microtubules into arrays through activities such as lattice severing, cross-linking, and sliding of microtubule pairs (Watanabe et al., 2005; Subramanian and Kapoor, 2012; Gardiner, 2013; Kapitein and Hoogenraad, 2015; Bodakuntla et al., 2019). For these reasons, tubulin heterodimers and MAPs have traditionally been the focal point in understanding the mechanisms that determine the assembly and overall architecture of an array. Microtubules themselves are largely viewed as passive building blocks in this process. A notable exception is the ‘tubulin code’, which is generated through the diversity in tubulin isotypes and post-translational modifications on the microtubule lattice. These chemical modifications of tubulin at the amino acid level can affect the inherent physical properties of the polymer and also regulate MAP activity (Alper et al., 2014; Kaul et al., 2014; Sirajuddin et al., 2014; Leo et al., 2015; Valenstein and Roll-Mecak, 2016; Bailey et al., 2015; Schwarz et al., 1998; Robison et al., 2016; Portran et al., 2017). An in-depth discussion of the tubulin code hypothesis can be found in several excellent reviews and will not be discussed further (Janke and Kneussel, 2010; Song and Brady, 2015; Gadadhar et al., 2017; Wloga et al., 2017; Janke and Magiera, 2020; Verhey and Gaertig, 2007). In recent years, an exciting concept that has emerged is that microtubules can encode information on the micron-scale to direct the self-organization of arrays. The features of the microtubule lattice that provide this information fall into two categories: (i) structural defects on the microtubule lattice such as missing tubulin subunits or changes in protofilament arrangement and (ii) size and relative arrangement of polymers, specified by parameters such as microtubule length, and the overlap length or contact angle between two adjacent polymers. Together, these micron-scale geometrical features along with the nanometer-scale tubulin code modulate the dynamics of individual polymers and regulate the activity of associated MAPs. In the following sections, we discuss how regulation by geometrical features of lattices is central to several processes involved in building a multi-microtubule array and determining its architecture. Overall, these mechanisms highlight the role of microtubules - as master architects directing their assembly and organization into cellular arrays.

• ‘Would you tell me, please, which way I ought to go from here?’

• ‘That depends a good deal on where you want to get to’

• – Alice’s Adventures in Wonderland, Lewis Carroll

Microtubules in cells are organized into specialized arrays. Upon close inspection, arrays are seen to contain recurring motifs or arrangements of microtubules A frequently observed motif is one where two polymers are coaligned. This can result in networks where adjacent microtubules are parallel to each other such as in ciliary axonemes and neuronal axons, anti-parallel as in the midzone of the mitotic spindle, or form arrays of mixed polarity such as plant cortical arrays (Subramanian and Kapoor, 2012). In some cases, these aligned arrays emerge from microtubules that are initially randomly oriented relative to each other. How are randomly oriented microtubules coaligned? Several studies highlight the importance of a geometrical parameter, the contact angle, as an important determinant in mechanisms promoting microtubule coalignment. For clarity, we define the contact angle as the angle between the lines joining the point where two microtubules encounter each other and their respective minus-ends (Figure 2).

Figure 2

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• ‘Oh, I'm not particular as to size, only one doesn't like changing so often, you know’

• – Alice’s Adventures in Wonderland, Lewis Carroll

An army of diverse Microtubule Associated Proteins MAPs serve as ‘builders’ to determine the size and shape of arrays through functions such as regulating microtubule dynamics, severing, cross-linking, and relative sliding between microtubules (van de Willige et al., 2016; Elliott and Shaw, 2018; Conkar and Firat-Karalar, 2021). The activity of MAPs, in turn, is found to be modulated by the geometrical parameters that specify array dimensions. On a single microtubule, the relevant parameter is polymer length. For pairs of microtubules, additional geometrical parameters that arise include the relative polarity, ‘overlap length’ over which two adjacent microtubules are cross-linked, the ‘overlap width’, which is the perpendicular distance between their lattices, and contact angle at their site of encounter. In the following section, we discuss the mechanisms by which these geometrical features regulate MAPs, and highlight how the interplay between geometrical parameters and MAP activity sculpt cellular arrays.

Part 1: Microtubule length as a regulator of microtubule dynamic instability

The lengths of individual microtubules are determined by protein regulators of microtubule dynamic instability (Howard and Hyman, 2007). Several studies have uncovered the existence of feedback loops between regulator activity and microtubule length. For example, in budding yeast astral and kinetochore microtubules, the rate of tubulin disassembly by the kinesin-5 motor Cin8p increases with microtubule length (Gardner et al., 2008). Similarly in vitro studies reveal that the rate of microtubule depolymerization by kinesin-8 family members Kip3p and Kif18A and the rate of microtubule polymerization by the budding yeast kinesin Kip2p are proportional to microtubule length (Figure 3(i); Gupta et al., 2006; Varga et al., 2006; Mayr et al., 2007; Varga et al., 2009; Hibbel et al., 2015).

Figure 3

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The microtubule length-dependent regulation of microtubule dynamics has been best studied in the context of Kip3p and explained by a mechanism called the ‘antenna model’ (Varga et al., 2006). Here, motor molecules land stochastically all along the lattice and step toward microtubule plus-ends. For highly processive motors with a slow dissociation rate from the microtubule end, such as in the case of Kip3p, the number of motors accumulated on the plus-ends is proportional to microtubule length (Figure 3(i)). Consequently, the rate of polymerization or depolymerization at the tip, which is mediated by motors, scales with microtubule length. An increase in depolymerization activity at the plus-end shortens microtubules, thereby providing negative feedback by reducing further motor accumulation and depolymerization rates. The antenna model thus illuminates the biophysical principles by which highly processive motors or motor complexes can alter microtubule dynamics in a microtubule length-dependent manner [(Subramanian et al., 2013; Leduc et al., 2012 Varga et al., 2006).].

• ‘I know who I was when I got up this morning, but I think I must have been changed several times since then.’

• – Alice’s Adventures in Wonderland, Lewis Carroll

In the crowded cellular volume, growing microtubules encounter physical obstacles such as other cytoskeletal arrays, cell membranes, and chromosomes, which can cause the filaments to undergo bending, buckling, or bundling (Odde et al., 1999; Waterman-Storer and Salmon, 1997). The stepping of motor proteins and the action of severing enzymes can also physically damage the lattice (Srayko et al., 2006; Dumont et al., 2015). As a result, both in vitro and in cells, microtubules display ‘lattice defects’, which are disruptions in the ordered arrangement of tubulin subunits in a polymer, extending over length scales ranging from 0.1 to 1 μm (Chrétien et al., 1992; Atherton et al., 2018). These defects manifest as vacancies in the lattice caused by missing tubulin heterodimers, sharp edges on the tubule due to abrupt shifts in protofilament number, or as regions of structural heterogeneity. Despite being localized to discrete sites along the microtubule lattice, defects have been shown to affect plus-end dynamics. For instance, a mismatch in protofilament number on the lattice can lead to an increase in catastrophe frequency (Rai et al., 2021). Sites of microtubule damage can also recruit MAPs whose activity changes the total density of microtubules in cellular arrays through the following known mechanisms.

Lattice repair increases microtubule lifetimes

Somewhat counterintuitively, studies in animal cells have shown that damage to microtubule lattices can result in an increase in the length and stability of microtubules (Aumeier et al., 2016). Several lines of experimental evidence indicate that sites of damage in the middle of a microtubule can incorporate GTP-tubulin dimers, as observed in live cells (Dimitrov et al., 2008; de Forges et al., 2016) and in microtubules polymerized in vitro (Schaedel et al., 2015; Schaedel et al., 2019; Rai et al., 2020). These ‘GTP-islands’ promote microtubule growth in two ways: (i) by serving as rescue points for re-initiation of growth during subsequent catastrophe events (Dimitrov et al., 2008; Tropini et al., 2012; Aumeier et al., 2016) and (ii) through the recruitment of growth-promoting MAPs that specifically recognize and bind to GTP-tubulin over GDP-tubulin, such as the rescue factors cytoplasmic linker protein of 170 kDa (CLIP-170) and CLIP-associating protein (CLASP) (Figure 4(i); Folker et al., 2005; de Forges et al., 2016; Aher et al., 2020). Consequently, as demonstrated by in vitro studies, sites of damage can result in microtubules that have significantly longer lifetimes, lengths, and rescue frequencies compared to microtubules undergoing stochastic polymerization and depolymerization (Aumeier et al., 2016). Thus, the repair of damaged lattices can infuse a fresh lease of life into a dynamic microtubule. This is advantageous in a crowded environment, where microtubules have a high probability of crossing over each other, which can result in lattice damage (de Forges et al., 2016). Repair at a site of damage increases the rescue frequency, resulting in an increase in the average length of polymers in the network. This further enhances the likelihood of microtubule crossovers, lattice damage, and rescue, thus increasing the overall density and lifetime of microtubules in the array.

Figure 4

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• ‘Begin at the beginning,’ the King said gravely, ‘and go on till you come to the end: then stop.’

• – Alice’s Adventures in Wonderland, Lewis Carroll

Microtubule arrays are dynamically assembled, remodeled, and disassembled over the lifetime of a cell. For example, the organization of the spindle constantly changes during cell division, plant cortical arrays are reoriented en masse to specify a new cell elongation axis in response to external stimuli, the polarities of microtubule bundles in neurons change as minor neurites grow into axons and dendrites, and the ciliary axoneme is built and disassembled each cell cycle (Sánchez and Dynlacht, 2016; van Beuningen and Hoogenraad, 2016). The assembly and remodeling of cellular arrays involves the integration of the various microtubule-regulated mechanisms discussed above such as microtubule nucleation, coalignment, MAP regulation, lattice severing, and repair. These processes do not occur in a discrete or sequential manner, but in fact occur concurrently, as illustrated by the two examples below.

Reorganization of plant cortical arrays

Stimulation of plant cells by blue light results in a dramatic reorientation of their cortical microtubules by ~90°, to specify a new cell elongation axis (Zandomeni and Schopfer, 1993; Paredez et al., 2006; Chen et al., 2016). In this section, we refer to the initial array as the longitudinal array, and the final arrangement, which is perpendicular to the original array, as the transverse array (Figure 5(i)). The microtubules of the longitudinal array play a critical role in directing array reorientation (Lindeboom et al., 2013). The process begins with γ-TuRC-mediated branching microtubule nucleation, from the microtubules in the longitudinal array. This results in new microtubules which grow at angles close to ±40° (hereafter referred to as diagonal microtubules) to the longitudinal array. Subsequent branching nucleation from diagonal microtubules generates a subset of new microtubules that grow perpendicular to the longitudinal array and are oriented correctly to form the transverse array. However, to establish an array that is reoriented by 90°, the selective amplification of transverse microtubules and destabilization of longitudinal and diagonal microtubules is required. For this, the severing enzyme katanin is specifically recruited to sites where two microtubules cross over (Zhang et al., 2013; Deinum et al., 2017; Wang et al., 2017). At these sites, the frequency of katanin severing is higher on the new microtubule compared to microtubules of the original array, although the precise mechanism for selective severing is currently unknown. Katanin severing of a new microtubule generates two microtubules, doubling the number of transverse and diagonal microtubules that continue to grow (Nakamura et al., 2018). How does this mechanism preferentially amplify transverse over diagonal microtubules? A new microtubule forms the maximum number of cross-overs, when the distance between the pre-existing microtubules that it encounters is the least. Basic geometry dictates that this condition is satisfied when the new microtubule grows perpendicular to the original array, compared to all other orientations, thereby selectively amplifying the transverse microtubules. Once the transverse array is established, there is a reduction in branching nucleation rates by mechanisms such as phototropin signaling, and a reduction in severing at cross-overs by mechanisms that remain unclear. This halts both generation and amplification of new microtubules, and a stable transverse array is e. Thus, the longitudinal array serves as a patterned template for the preferential amplification of microtubules that grow at right angles to it, to form the transverse array (Figure 5(i)).

Figure 5

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Conclusion

The self-organization of microtubules into specialized micron-sized arrays has been extensively studied from the perspective of motor and non-motor MAPs that shape arrays. It is now evident that microtubules themselves provide cues to regulate MAP activity, both at the nanometer scale through a tubulin code and at the micron scale via geometrical features of the lattice. The interplay between microtubules and MAP activity directs the self-organization process, by specifying microtubule density, array architecture, and stability (Figure 6). This ‘DIY’ (Do It Yourself) approach adopted by microtubules toward their organization has far-reaching consequences in the cell. First, members of the different cytoskeletal families are known to engage in extensive crosstalk. (Dogterom and Koenderink, 2019; Logan and Menko, 2019; Seetharaman and Etienne-Manneville, 2020). For example, microtubules have been shown to regulate the nucleation, polymerization, and dynamics of the actin network in migrating cells, neuronal growth cones, and dendritic spines. In dividing cells, the position and geometry of the acto-myosin contractile ring at the cell center are determined by astral and spindle midzone microtubules (Pollard and O'Shaughnessy, 2019). Further, microtubule-based transport is also important for the assembly of intermediate filaments (Hookway et al., 2015). Second, microtubule networks are important in creating and maintaining biochemical concentration gradients in the cytosol. For instance, spindle microtubules are known to play a role in creating spatial gradients of Aurora B kinase activity and components of the Ran-GTP pathway (Noujaim et al., 2014; Oh et al., 2016). Thus, beyond directing the assembly of microtubules into arrays, micron-sized features of microtubule networks contribute to the larger-scale spatial organization of the eukaryotic cell by directing the organization of other cytoskeletal networks and signaling gradients.

Figure 6

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Polymeric proteins that self-organize into micron-sized arrays are not exclusive to eukaryotic cells, but are found across all kingdoms of life. In bacteria and archaea, homologs of tubulin, actin, and intermediate filaments form filamentous assemblies, with the assistance of conserved proteins (Busiek and Margolin, 2015; Fink et al., 2016; Stairs and Ettema, 2020). Future studies will reveal if the mechanisms invoking the regulation of microtubule array assembly by geometrical features of microtubules could have emerged in prokaryotic ancestors. In addition to the assembly of protein-based structures, these principles may also apply to the self-organization of nucleic acids and other cellular polymeric arrays.

• Multiple nucleation pathways co-exist in cells in addition to microtubule-templated microtubule nucleation, each yielding new filaments that grow at specific orientation and polarity with respect to their organizing centers. For instance, centrosome-nucleated microtubules grow out radially, golgi-nucleated microtubules are tangential to the golgi, and nuclear membrane-associated microtubules form longitudinal arrays (Miller et al., 2009; Lüders and Stearns, 2007; Musa et al., 2003). Whether there are specialized geometrical cues that dictate the organization of these arrays relative to pre-existing microtubule networks presents an exciting research direction.

• Several mechanisms facilitate the close packing of microtubules to form bundles, ranging from those that occur in the absence of all MAPs to others which utilize cross-linking proteins of varying lengths (Peterman and Scholey, 2009; Needleman et al., 2004b). Thus far, our understanding of the self-organization and function of microtubule bundles have largely come from studies on coaligned microtubules cross-linked by members of the Ase1p/PRC1 family and a few cross-linking motor proteins of the kinesin family. Physical mechanisms such as osmotic forces induced by macromolecules and formation of liquid crystalline domains also result in the bundling of microtubules (Hitt et al., 1990; Needleman et al., 2004a; Guo et al., 2008). Remarkably, the shape of microtubule bundles formed by these mechanisms and the motility of motor proteins within them are found to be different when compared to bundles formed with protein cross-linkers (Edozie et al., 2019; Conway et al., 2014). Whether the close packing of microtubules in the absence of all MAPs generates geometrical cues similar to those observed with protein cross-linkers is still under-explored. Extending this line of research to diverse protein and non-protein cross-linkers will expand our understanding of geometrical parameters in array organization.

• The role of geometrical parameters in the organization of microtubules into radial, polar arrays such as asters remains poorly understood (Needleman and Dogic, 2017; Rickman et al., 2019). Intriguingly, in vitro studies show that motor proteins belonging to the kinesin-5 or kinesin-14 families can promote either nematic (linear bundles) or polar (radial arrays) organization of microtubules, depending on motor velocity and number, soluble tubulin concentration, microtubule number, and growth rates (Roostalu et al., 2018; Norris et al., 2018). Whether geometric features of microtubules direct the formation of polar arrays by kinesin and dynein motors (Tan et al., 2018; Khetan et al., 2021), and mediate the switch between nematic and polar self-organization remains an open question.

• In addition to overlap length and intersection angles, a fundamental geometrical parameter that defines the relative placement of two microtubules within a close-packed network is the overlap width or the spacing between their lattices. The observation that cross-linking motors and MAPs are characterized by a range of molecular dimensions suggests that the overlap width parameter could (i) regulate selective access of MAPs to the microtubules in an array, which in turn can modulate bundle dynamics and structure (Conway et al., 2014), (ii) determine the 2D and 3D organization of polymers in an array, by varying the nearest-neighbor distances (Nixon et al., 2015; Needleman et al., 2004b). Reconstitution studies that systematically vary the overlap width are likely to provide more insights. The effect of overlap width and the size of the molecular cross-linkers is particularly interesting in the context of asters where the overlap width would vary along the lattice in a radial assembly of microtubules.

• Lattice defects such as bends and breaks in the microtubule wall have been shown in vitro to serve as entry points into the microtubule lumen for the enzyme αTAT1 where it acetylates tubulin on a residue facing the lumen (Coombes et al., 2016). This mechanism raises the exciting possibility that regions of missing tubulin subunits could serve as ‘sieves’, allowing for selective lumen entry of proteins based on size. In addition, a break in the ordered arrangement of tubulin subunits could also hinder motor motility. Precise measurements of the dimensions of lattice defects, including the number of tubulin dimers they span, number of protofilaments they extend over, and their effect on microtubule stability, motor motility, cross-linking, and severing probability will provide insights into how microtubule defects regulate microtubule organization.

• Microtubules serve as platforms to couple the activity of associated motors and non-motor MAPs. The best understood coupling mechanisms arise from short-range interactions such as steric effects and cooperativity or crowding (Vilfan et al., 2001; Leduc et al., 2012). Can the linear structure of microtubules mediate the coupling of MAP molecules over distances in the micron length scale? Prior literature raises the intriguing possibility that microtubules can act as a micron-scale coupling medium to influence microtubule-binding kinetics of MAPs and motile propertiesof motors (Muto et al., 2005; Wijeratne et al., 2020). For example, microtubule-bound kinesin-1 motors have been shown to increase the axial pitch of the microtubule lattice, an alteration that is thought to promote the cooperative binding of other kinesin-1 molecules (Peet et al., 2018; Shima et al., 2018). Similarly, the straightening of tips of protofilaments by EB1 is proposed to mediate the synergistic increase in microtubule growth rates in the presence of Xenopus microtubule-associated protein (XMAP215) and EB1 (Zanic et al., 2013). In addition to MAP binding and activity, long-range effects on the microtubule lattice can also modulate the dynamics of the polymer itself. For instance, the presence of a lattice defect site corresponding to protofilament number mismatch has been shown to affect the dynamics of the polymer’s plus-end which can be several microns away (Rai et al., 2021). Elucidating conditions for micron-scale coupling and uncovering new mechanisms by which the microtubule lattice facilitates long-range communication and allosteric regulation of MAPs promises to be an exciting new research direction.

1. 1. Aher A
   2. Rai D
   3. Schaedel L
   4. Gaillard J
   5. John K
   6. Liu Q
   7. Altelaar M
   8. Blanchoin L
   9. Thery M
   10. Akhmanova A

   (2020) CLASP mediates microtubule repair by restricting lattice damage and regulating tubulin incorporation

   Current Biology 30:2175–2183.

   https://doi.org/10.1016/j.cub.2020.03.070

   • Google Scholar
2. 1. Alfaro-Aco R
   2. Thawani A
   3. Petry S

   (2020) Biochemical reconstitution of branching microtubule nucleation

   eLife 9:e49797.

   https://doi.org/10.7554/eLife.49797

   • PubMed
   • Google Scholar
3. 1. Alper JD
   2. Decker F
   3. Agana B
   4. Howard J

   (2014) The motility of axonemal dynein is regulated by the tubulin code

   Biophysical Journal 107:2872–2880.

   https://doi.org/10.1016/j.bpj.2014.10.061

   • PubMed
   • Google Scholar
4. 1. Atherton J
   2. Stouffer M
   3. Francis F
   4. Moores CA

   (2018) Microtubule architecture in vitro and in cells revealed by cryo-electron tomography

   Acta Crystallographica Section D Structural Biology 74:572–584.

   https://doi.org/10.1107/S2059798318001948

   • PubMed
   • Google Scholar
5. 1. Aumeier C
   2. Schaedel L
   3. Gaillard J
   4. John K
   5. Blanchoin L
   6. Théry M

   (2016) Self-repair promotes microtubule rescue

   Nature Cell Biology 18:1054–1064.

   https://doi.org/10.1038/ncb3406

   • PubMed
   • Google Scholar
6. 1. Bacallao R
   2. Antony C
   3. Dotti C
   4. Karsenti E
   5. Stelzer EH
   6. Simons K

   (1989) The subcellular organization of Madin-Darby canine kidney cells during the formation of a polarized epithelium

   Journal of Cell Biology 109:2817–2832.

   https://doi.org/10.1083/jcb.109.6.2817

   • PubMed
   • Google Scholar
7. 1. Bailey ME
   2. Sackett DL
   3. Ross JL

   (2015) Katanin severing and binding microtubules are inhibited by tubulin carboxy tails

   Biophysical Journal 109:2546–2561.

   https://doi.org/10.1016/j.bpj.2015.11.011

   • PubMed
   • Google Scholar
8. 1. Basnet N
   2. Nedozralova H
   3. Crevenna AH
   4. Bodakuntla S
   5. Schlichthaerle T
   6. Taschner M
   7. Cardone G
   8. Janke C
   9. Jungmann R
   10. Magiera MM
   11. Biertümpfel C
   12. Mizuno N

   (2018) Direct induction of microtubule branching by microtubule nucleation factor SSNA1

   Nature Cell Biology 20:1172–1180.

   https://doi.org/10.1038/s41556-018-0199-8

   • PubMed
   • Google Scholar
9. 1. Bieling P
   2. Telley IA
   3. Surrey T

   (2010) A minimal midzone protein module controls formation and length of antiparallel microtubule overlaps

   Cell 142:420–432.

   https://doi.org/10.1016/j.cell.2010.06.033

   • PubMed
   • Google Scholar
10. 1. Bodakuntla S
    2. Jijumon AS
    3. Villablanca C
    4. Gonzalez-Billault C
    5. Janke C

    (2019) Microtubule-Associated proteins: structuring the cytoskeleton

    Trends in Cell Biology 29:804–819.

    https://doi.org/10.1016/j.tcb.2019.07.004

    • PubMed
    • Google Scholar
11. 1. Braun M
    2. Lansky Z
    3. Fink G
    4. Ruhnow F
    5. Diez S
    6. Janson ME

    (2011) Adaptive braking by Ase1 prevents overlapping microtubules from sliding completely apart

    Nature Cell Biology 13:1259–1264.

    https://doi.org/10.1038/ncb2323

    • PubMed
    • Google Scholar
12. 1. Braun M
    2. Lansky Z
    3. Hilitski F
    4. Dogic Z
    5. Diez S

    (2016) Entropic forces drive contraction of cytoskeletal networks

    BioEssays 38:474–481.

    https://doi.org/10.1002/bies.201500183

    • PubMed
    • Google Scholar
13. 1. Braun M
    2. Lansky Z
    3. Szuba A
    4. Schwarz FW
    5. Mitra A
    6. Gao M
    7. Lüdecke A
    8. Ten Wolde PR
    9. Diez S

    (2017) Changes in microtubule overlap length regulate kinesin-14-driven microtubule sliding

    Nature Chemical Biology 13:1245–1252.

    https://doi.org/10.1038/nchembio.2495

    • PubMed
    • Google Scholar
14. 1. Busiek KK
    2. Margolin W

    (2015) Bacterial actin and tubulin homologs in cell growth and division

    Current Biology 25:R243–R254.

    https://doi.org/10.1016/j.cub.2015.01.030

    • PubMed
    • Google Scholar
15. 1. Chen Y
    2. Rolls MM
    3. Hancock WO

    (2014) An EB1-kinesin complex is sufficient to steer microtubule growth in vitro

    Current Biology 24:316–321.

    https://doi.org/10.1016/j.cub.2013.11.024

    • PubMed
    • Google Scholar
16. 1. Chen X
    2. Wu S
    3. Liu Z
    4. Friml J

    (2016) Environmental and endogenous control of cortical microtubule orientation

    Trends in Cell Biology 26:409–419.

    https://doi.org/10.1016/j.tcb.2016.02.003

    • PubMed
    • Google Scholar
17. 1. Chi Z
    2. Ambrose C

    (2016) Microtubule encounter-based catastrophe in Arabidopsis cortical microtubule arrays

    BMC Plant Biology 16:18.

    https://doi.org/10.1186/s12870-016-0703-x

    • PubMed
    • Google Scholar
18. 1. Chrétien D
    2. Metoz F
    3. Verde F
    4. Karsenti E
    5. Wade RH

    (1992) Lattice defects in microtubules: protofilament numbers vary within individual microtubules

    Journal of Cell Biology 117:1031–1040.

    https://doi.org/10.1083/jcb.117.5.1031

    • Google Scholar
19. 1. Cleveland DW

    (1988) Autoregulated instability of tubulin mRNAs: a novel eukaryotic regulatory mechanism

    Trends in Biochemical Sciences 13:339–343.

    https://doi.org/10.1016/0968-0004(88)90103-X

    • PubMed
    • Google Scholar
20. 1. Conkar D
    2. Firat-Karalar EN

    (2021) Microtubule-associated proteins and emerging links to primary cilium structure, assembly, maintenance, and disassembly

    The FEBS Journal 288:786–798.

    https://doi.org/10.1111/febs.15473

    • PubMed
    • Google Scholar
21. 1. Consolati T
    2. Locke J
    3. Roostalu J
    4. Chen ZA
    5. Gannon J
    6. Asthana J
    7. Lim WM
    8. Martino F
    9. Cvetkovic MA
    10. Rappsilber J
    11. Costa A
    12. Surrey T

    (2020) Microtubule nucleation properties of single human γturcs explained by their Cryo-EM structure

    Developmental Cell 53:603–617.

    https://doi.org/10.1016/j.devcel.2020.04.019

    • Google Scholar
22. 1. Conway L
    2. Gramlich MW
    3. Ali Tabei SM
    4. Ross JL

    (2014) Microtubule orientation and spacing within bundles is critical for long-range kinesin-1 motility

    Cytoskeleton 71:595–610.

    https://doi.org/10.1002/cm.21197

    • PubMed
    • Google Scholar
23. 1. Coombes C
    2. Yamamoto A
    3. McClellan M
    4. Reid TA
    5. Plooster M
    6. Luxton GW
    7. Alper J
    8. Howard J
    9. Gardner MK

    (2016) Mechanism of microtubule lumen entry for the α-tubulin acetyltransferase enzyme αtat1

    PNAS 113:E7176–E7184.

    https://doi.org/10.1073/pnas.1605397113

    • PubMed
    • Google Scholar
24. 1. Cunha-Ferreira I
    2. Chazeau A
    3. Buijs RR
    4. Stucchi R
    5. Will L
    6. Pan X
    7. Adolfs Y
    8. van der Meer C
    9. Wolthuis JC
    10. Kahn OI
    11. Schätzle P
    12. Altelaar M
    13. Pasterkamp RJ
    14. Kapitein LC
    15. Hoogenraad CC

    (2018) The HAUS complex is a key regulator of Non-centrosomal microtubule organization during neuronal development

    Cell Reports 24:791–800.

    https://doi.org/10.1016/j.celrep.2018.06.093

    • PubMed
    • Google Scholar
25. 1. David AF
    2. Roudot P
    3. Legant WR
    4. Betzig E
    5. Danuser G
    6. Gerlich DW

    (2019) Augmin accumulation on long-lived microtubules drives amplification and kinetochore-directed growth

    Journal of Cell Biology 218:2150–2168.

    https://doi.org/10.1083/jcb.201805044

    • PubMed
    • Google Scholar
26. 1. Davis LJ
    2. Odde DJ
    3. Block SM
    4. Gross SP

    (2002) The importance of lattice defects in katanin-mediated microtubule severing in vitro

    Biophysical Journal 82:2916–2927.

    https://doi.org/10.1016/S0006-3495(02)75632-4

    • PubMed
    • Google Scholar
27. 1. de Forges H
    2. Pilon A
    3. Cantaloube I
    4. Pallandre A
    5. Haghiri-Gosnet AM
    6. Perez F
    7. Poüs C

    (2016) Localized mechanical stress promotes microtubule rescue

    Current Biology 26:3399–3406.

    https://doi.org/10.1016/j.cub.2016.10.048

    • PubMed
    • Google Scholar
28. 1. Deinum EE
    2. Tindemans SH
    3. Lindeboom JJ
    4. Mulder BM

    (2017) How selective severing by Katanin promotes order in the plant cortical microtubule array

    PNAS 114:6942–6947.

    https://doi.org/10.1073/pnas.1702650114

    • PubMed
    • Google Scholar
29. 1. Díaz-Valencia JD
    2. Morelli MM
    3. Bailey M
    4. Zhang D
    5. Sharp DJ
    6. Ross JL

    (2011) Drosophila katanin-60 depolymerizes and severs at Microtubule defects

    Biophysical Journal 100:2440–2449.

    https://doi.org/10.1016/j.bpj.2011.03.062

    • PubMed
    • Google Scholar
30. 1. Dimitrov A
    2. Quesnoit M
    3. Moutel S
    4. Cantaloube I
    5. Poüs C
    6. Perez F

    (2008) Detection of GTP-tubulin conformation in vivo reveals a role for GTP remnants in microtubule rescues

    Science 322:1353–1356.

    https://doi.org/10.1126/science.1165401

    • PubMed
    • Google Scholar
31. 1. Dixit R
    2. Chang E
    3. Cyr R

    (2006) Establishment of polarity during organization of the acentrosomal plant cortical microtubule array

    Molecular Biology of the Cell 17:1298–1305.

    https://doi.org/10.1091/mbc.e05-09-0864

    • PubMed
    • Google Scholar
32. 1. Dixit R
    2. Cyr R

    (2004) Encounters between dynamic cortical microtubules promote ordering of the cortical array through angle-dependent modifications of microtubule behavior

    The Plant Cell 16:3274–3284.

    https://doi.org/10.1105/tpc.104.026930

    • PubMed
    • Google Scholar
33. 1. Dogterom M
    2. Koenderink GH

    (2019) Actin-microtubule crosstalk in cell biology

    Nature Reviews Molecular Cell Biology 20:38–54.

    https://doi.org/10.1038/s41580-018-0067-1

    • PubMed
    • Google Scholar
34. 1. Doodhi H
    2. Katrukha EA
    3. Kapitein LC
    4. Akhmanova A

    (2014) Mechanical and geometrical constraints control kinesin-based microtubule guidance

    Current Biology 24:322–328.

    https://doi.org/10.1016/j.cub.2014.01.005

    • PubMed
    • Google Scholar
35. 1. Dumont EL
    2. Do C
    3. Hess H

    (2015) Molecular wear of microtubules propelled by surface-adhered kinesins

    Nature Nanotechnology 10:166–169.

    https://doi.org/10.1038/nnano.2014.334

    • PubMed
    • Google Scholar
36. 1. Edozie B
    2. Sahu S
    3. Pitta M
    4. Englert A
    5. do Rosario CF
    6. Ross JL

    (2019) Self-organization of spindle-like microtubule structures

    Soft Matter 15:4797–4807.

    https://doi.org/10.1039/C8SM01835A

    • PubMed
    • Google Scholar
37. 1. Eggert US
    2. Mitchison TJ
    3. Field CM

    (2006) Animal cytokinesis: from parts list to mechanisms

    Annual Review of Biochemistry 75:543–566.

    https://doi.org/10.1146/annurev.biochem.74.082803.133425

    • PubMed
    • Google Scholar
38. 1. Ehrhardt DW
    2. Shaw SL

    (2006) Microtubule dynamics and organization in the plant cortical array

    Annual Review of Plant Biology 57:859–875.

    https://doi.org/10.1146/annurev.arplant.57.032905.105329

    • PubMed
    • Google Scholar
39. 1. Elliott A
    2. Shaw SL

    (2018) Update: plant cortical microtubule arrays

    Plant Physiology 176:94–105.

    https://doi.org/10.1104/pp.17.01329

    • PubMed
    • Google Scholar
40. 1. Fink G
    2. Szewczak-Harris A
    3. Löwe J

    (2016) SnapShot: the bacterial cytoskeleton

    Cell 166:522.

    https://doi.org/10.1016/j.cell.2016.06.057

    • PubMed
    • Google Scholar
41. 1. Folker ES
    2. Baker BM
    3. Goodson HV

    (2005) Interactions between CLIP-170, tubulin, and microtubules: implications for the mechanism of Clip-170 plus-end tracking behavior

    Molecular Biology of the Cell 16:5373–5384.

    https://doi.org/10.1091/mbc.e04-12-1106

    • PubMed
    • Google Scholar
42. 1. Forth S
    2. Kapoor TM

    (2017) The mechanics of microtubule networks in cell division

    Journal of Cell Biology 216:1525–1531.

    https://doi.org/10.1083/jcb.201612064

    • PubMed
    • Google Scholar
43. 1. Gadadhar S
    2. Bodakuntla S
    3. Natarajan K
    4. Janke C

    (2017) The tubulin code at a glance

    Journal of Cell Science 130:1347–1353.

    https://doi.org/10.1242/jcs.199471

    • PubMed
    • Google Scholar
44. 1. Gardiner J

    (2013) The evolution and diversification of plant microtubule-associated proteins

    The Plant Journal 75:219–229.

    https://doi.org/10.1111/tpj.12189

    • PubMed
    • Google Scholar
45. 1. Gardner MK
    2. Bouck DC
    3. Paliulis LV
    4. Meehl JB
    5. O'Toole ET
    6. Haase J
    7. Soubry A
    8. Joglekar AP
    9. Winey M
    10. Salmon ED
    11. Bloom K
    12. Odde DJ

    (2008) Chromosome congression by Kinesin-5 motor-mediated disassembly of longer kinetochore microtubules

    Cell 135:894–906.

    https://doi.org/10.1016/j.cell.2008.09.046

    • PubMed
    • Google Scholar
46. 1. Gaska I
    2. Armstrong ME
    3. Alfieri A
    4. Forth S

    (2020) The mitotic crosslinking protein PRC1 acts like a mechanical dashpot to resist microtubule sliding

    Developmental Cell 54:367–378.

    https://doi.org/10.1016/j.devcel.2020.06.017

    • Google Scholar
47. 1. Gerson-Gurwitz A
    2. Thiede C
    3. Movshovich N
    4. Fridman V
    5. Podolskaya M
    6. Danieli T
    7. Lakämper S
    8. Klopfenstein DR
    9. Schmidt CF
    10. Gheber L

    (2011) Directionality of individual kinesin-5 Cin8 motors is modulated by loop 8, ionic strength and microtubule geometry

    The EMBO Journal 30:4942–4954.

    https://doi.org/10.1038/emboj.2011.403

    • PubMed
    • Google Scholar
48. 1. Glotzer M

    (2009) The 3ms of central spindle assembly: microtubules, motors and MAPs

    Nature Reviews Molecular Cell Biology 10:9–20.

    https://doi.org/10.1038/nrm2609

    • PubMed
    • Google Scholar
49. 1. Goshima G
    2. Mayer M
    3. Zhang N
    4. Stuurman N
    5. Vale RD

    (2008) Augmin: a protein complex required for centrosome-independent microtubule generation within the spindle

    Journal of Cell Biology 181:421–429.

    https://doi.org/10.1083/jcb.200711053

    • PubMed
    • Google Scholar
50. 1. Grover R
    2. Fischer J
    3. Schwarz FW
    4. Walter WJ
    5. Schwille P
    6. Diez S

    (2016) Transport efficiency of membrane-anchored kinesin-1 motors depends on motor density and diffusivity

    PNAS 113:E7185–E7193.

    https://doi.org/10.1073/pnas.1611398113

    • PubMed
    • Google Scholar
51. 1. Guo Y
    2. Liu Y
    3. Oldenbourg R
    4. Tang JX
    5. Valles JM

    (2008) Effects of osmotic force and torque on microtubule bundling and pattern formation

    Physical Review E 78:041910.

    https://doi.org/10.1103/PhysRevE.78.041910

    • Google Scholar
52. 1. Gupta ML
    2. Carvalho P
    3. Roof DM
    4. Pellman D

    (2006) Plus end-specific depolymerase activity of Kip3, a kinesin-8 protein, explains its role in positioning the yeast mitotic spindle

    Nature Cell Biology 8:913–923.

    https://doi.org/10.1038/ncb1457

    • PubMed
    • Google Scholar
53. 1. Hannabuss J
    2. Lera-Ramirez M
    3. Cade NI
    4. Fourniol FJ
    5. Nédélec F
    6. Surrey T

    (2019) Self-Organization of minimal anaphase spindle midzone bundles

    Current Biology 29:2120–2130.

    https://doi.org/10.1016/j.cub.2019.05.049

    • Google Scholar
54. 1. Hibbel A
    2. Bogdanova A
    3. Mahamdeh M
    4. Jannasch A
    5. Storch M
    6. Schäffer E
    7. Liakopoulos D
    8. Howard J

    (2015) Kinesin Kip2 enhances microtubule growth in vitro through length-dependent feedback on polymerization and catastrophe

    eLife 4:e10542.

    https://doi.org/10.7554/eLife.10542

    • PubMed
    • Google Scholar
55. 1. Hitt AL
    2. Cross AR
    3. Williams RC

    (1990) Microtubule solutions display nematic liquid crystalline structure

    Journal of Biological Chemistry 265:1639–1647.

    https://doi.org/10.1016/S0021-9258(19)40064-1

    • PubMed
    • Google Scholar
56. 1. Hookway C
    2. Ding L
    3. Davidson MW
    4. Rappoport JZ
    5. Danuser G
    6. Gelfand VI

    (2015) Microtubule-dependent transport and dynamics of vimentin intermediate filaments

    Molecular Biology of the Cell 26:1675–1686.

    https://doi.org/10.1091/mbc.E14-09-1398

    • PubMed
    • Google Scholar
57. 1. Howard J
    2. Hudspeth AJ
    3. Vale RD

    (1989) Movement of microtubules by single kinesin molecules

    Nature 342:154–158.

    https://doi.org/10.1038/342154a0

    • PubMed
    • Google Scholar
58. 1. Howard J
    2. Hyman AA

    (2007) Microtubule polymerases and depolymerases

    Current Opinion in Cell Biology 19:31–35.

    https://doi.org/10.1016/j.ceb.2006.12.009

    • PubMed
    • Google Scholar
59. 1. Hunt AJ
    2. Gittes F
    3. Howard J

    (1994) The force exerted by a single kinesin molecule against a viscous load

    Biophysical Journal 67:766–781.

    https://doi.org/10.1016/S0006-3495(94)80537-5

    • PubMed
    • Google Scholar
60. 1. Janke C
    2. Kneussel M

    (2010) Tubulin post-translational modifications: encoding functions on the neuronal microtubule cytoskeleton

    Trends in Neurosciences 33:362–372.

    https://doi.org/10.1016/j.tins.2010.05.001

    • PubMed
    • Google Scholar
61. 1. Janke C
    2. Magiera MM

    (2020) The tubulin code and its role in controlling microtubule properties and functions

    Nature Reviews Molecular Cell Biology 21:307–326.

    https://doi.org/10.1038/s41580-020-0214-3

    • PubMed
    • Google Scholar
62. 1. Janson ME
    2. de Dood ME
    3. Dogterom M

    (2003) Dynamic instability of microtubules is regulated by force

    Journal of Cell Biology 161:1029–1034.

    https://doi.org/10.1083/jcb.200301147

    • PubMed
    • Google Scholar
63. 1. Janson ME
    2. Setty TG
    3. Paoletti A
    4. Tran PT

    (2005) Efficient formation of bipolar microtubule bundles requires microtubule-bound gamma-tubulin complexes

    Journal of Cell Biology 169:297–308.

    https://doi.org/10.1083/jcb.200410119

    • PubMed
    • Google Scholar
64. 1. Janson ME
    2. Loughlin R
    3. Loïodice I
    4. Fu C
    5. Brunner D
    6. Nédélec FJ
    7. Tran PT

    (2007) Crosslinkers and motors organize dynamic microtubules to form stable bipolar arrays in fission yeast

    Cell 128:357–368.

    https://doi.org/10.1016/j.cell.2006.12.030

    • PubMed
    • Google Scholar
65. 1. Kapitein LC
    2. Kwok BH
    3. Weinger JS
    4. Schmidt CF
    5. Kapoor TM
    6. Peterman EJ

    (2008) Microtubule cross-linking triggers the directional motility of kinesin-5

    Journal of Cell Biology 182:421–428.

    https://doi.org/10.1083/jcb.200801145

    • PubMed
    • Google Scholar
66. 1. Kapitein LC
    2. Hoogenraad CC

    (2015) Building the neuronal microtubule cytoskeleton

    Neuron 87:492–506.

    https://doi.org/10.1016/j.neuron.2015.05.046

    • PubMed
    • Google Scholar
67. 1. Karsenti E

    (2008) Self-organization in cell biology: a brief history

    Nature Reviews Molecular Cell Biology 9:255–262.

    https://doi.org/10.1038/nrm2357

    • PubMed
    • Google Scholar
68. 1. Kaul N
    2. Soppina V
    3. Verhey KJ

    (2014) Effects of α-tubulin K40 acetylation and detyrosination on kinesin-1 motility in a purified system

    Biophysical Journal 106:2636–2643.

    https://doi.org/10.1016/j.bpj.2014.05.008

    • PubMed
    • Google Scholar
69. 1. Kelliher MT
    2. Saunders HA
    3. Wildonger J

    (2019) Microtubule control of functional architecture in neurons

    Current Opinion in Neurobiology 57:39–45.

    https://doi.org/10.1016/j.conb.2019.01.003

    • PubMed
    • Google Scholar
70. 1. Khetan N
    2. Pruliere G
    3. Hebras C
    4. Chenevert J
    5. Athale CA

    (2021) Self-organized optimal packing of kinesin-5-driven microtubule asters scales with cell size

    Journal of Cell Science 134:jcs257543.

    https://doi.org/10.1242/jcs.257543

    • PubMed
    • Google Scholar
71. 1. King MR
    2. Petry S

    (2020) Phase separation of TPX2 enhances and spatially coordinates microtubule nucleation

    Nature Communications 11:270.

    https://doi.org/10.1038/s41467-019-14087-0

    • PubMed
    • Google Scholar
72. 1. Kollman JM
    2. Zelter A
    3. Muller EG
    4. Fox B
    5. Rice LM
    6. Davis TN
    7. Agard DA

    (2008) The structure of the gamma-tubulin small complex: implications of its architecture and flexibility for microtubule nucleation

    Molecular Biology of the Cell 19:207–215.

    https://doi.org/10.1091/mbc.e07-09-0879

    • PubMed
    • Google Scholar
73. 1. Kong Z
    2. Hotta T
    3. Lee YR
    4. Horio T
    5. Liu B

    (2010) The {gamma}-tubulin complex protein GCP4 is required for organizing functional microtubule arrays in Arabidopsis thaliana

    The Plant Cell 22:191–204.

    https://doi.org/10.1105/tpc.109.071191

    • PubMed
    • Google Scholar
74. 1. Krüger LK
    2. Sanchez JL
    3. Paoletti A
    4. Tran PT

    (2019) Kinesin-6 regulates cell-size-dependent spindle elongation velocity to keep mitosis duration constant in fission yeast

    eLife 8:e42182.

    https://doi.org/10.7554/eLife.42182

    • PubMed
    • Google Scholar
75. 1. Kuo YW
    2. Trottier O
    3. Mahamdeh M
    4. Howard J

    (2019) Spastin is a dual-function enzyme that severs microtubules and promotes their regrowth to increase the number and mass of microtubules

    PNAS 116:5533–5541.

    https://doi.org/10.1073/pnas.1818824116

    • PubMed
    • Google Scholar
76. 1. Leary A
    2. Sim S
    3. Nazarova E
    4. Shulist K
    5. Genthial R
    6. Yang SK
    7. Bui KH
    8. Francois P
    9. Vogel J

    (2019) Successive Kinesin-5 microtubule crosslinking and sliding promote fast, irreversible formation of a stereotyped bipolar spindle

    Current Biology 29:3825–3837.

    https://doi.org/10.1016/j.cub.2019.09.030

    • Google Scholar
77. 1. Leduc C
    2. Padberg-Gehle K
    3. Varga V
    4. Helbing D
    5. Diez S
    6. Howard J

    (2012) Molecular crowding creates traffic jams of kinesin motors on microtubules

    PNAS 109:6100–6105.

    https://doi.org/10.1073/pnas.1107281109

    • PubMed
    • Google Scholar
78. 1. Lee YJ
    2. Liu B

    (2019) Microtubule nucleation for the assembly of acentrosomal microtubule arrays in plant cells

    New Phytologist 222:1705–1718.

    https://doi.org/10.1111/nph.15705

    • PubMed
    • Google Scholar
79. 1. Leo L
    2. Yu W
    3. D'Rozario M
    4. Waddell EA
    5. Marenda DR
    6. Baird MA
    7. Davidson MW
    8. Zhou B
    9. Wu B
    10. Baker L
    11. Sharp DJ
    12. Baas PW

    (2015) Vertebrate fidgetin restrains axonal growth by severing labile domains of microtubules

    Cell Reports 12:1723–1730.

    https://doi.org/10.1016/j.celrep.2015.08.017

    • PubMed
    • Google Scholar
80. 1. Lera-Ramirez M
    2. Nédélec FJ

    (2019) Theory of antiparallel microtubule overlap stabilization by motors and diffusible crosslinkers

    Cytoskeleton 76:600–610.

    https://doi.org/10.1002/cm.21574

    • PubMed
    • Google Scholar
81. 1. Lin Z
    2. Gasic I
    3. Chandrasekaran V
    4. Peters N
    5. Shao S
    6. Mitchison TJ
    7. Hegde RS

    (2020) TTC5 mediates autoregulation of tubulin via mRNA degradation

    Science 367:100–104.

    https://doi.org/10.1126/science.aaz4352

    • PubMed
    • Google Scholar
82. 1. Lindeboom JJ
    2. Nakamura M
    3. Hibbel A
    4. Shundyak K
    5. Gutierrez R
    6. Ketelaar T
    7. Emons AM
    8. Mulder BM
    9. Kirik V
    10. Ehrhardt DW

    (2013) A mechanism for reorientation of cortical microtubule arrays driven by microtubule severing

    Science 342:1245533.

    https://doi.org/10.1126/science.1245533

    • PubMed
    • Google Scholar
83. 1. Liu T
    2. Tian J
    3. Wang G
    4. Yu Y
    5. Wang C
    6. Ma Y
    7. Zhang X
    8. Xia G
    9. Liu B
    10. Kong Z

    (2014) Augmin triggers microtubule-dependent microtubule nucleation in interphase plant cells

    Current Biology 24:2708–2713.

    https://doi.org/10.1016/j.cub.2014.09.053

    • PubMed
    • Google Scholar
84. 1. Liu P
    2. Zupa E
    3. Neuner A
    4. Böhler A
    5. Loerke J
    6. Flemming D
    7. Ruppert T
    8. Rudack T
    9. Peter C
    10. Spahn C
    11. Gruss OJ
    12. Pfeffer S
    13. Schiebel E

    (2020) Insights into the assembly and activation of the microtubule nucleator γ-TuRC

    Nature 578:467–471.

    https://doi.org/10.1038/s41586-019-1896-6

    • PubMed
    • Google Scholar
85. 1. Liu P
    2. Würtz M
    3. Zupa E
    4. Pfeffer S
    5. Schiebel E

    (2021) Microtubule nucleation: the waltz between γ-tubulin ring complex and associated proteins

    Current Opinion in Cell Biology 68:124–131.

    https://doi.org/10.1016/j.ceb.2020.10.004

    • PubMed
    • Google Scholar
86. 1. Logan CM
    2. Menko AS

    (2019) Microtubules: evolving roles and critical cellular interactions

    Experimental Biology and Medicine 244:1240–1254.

    https://doi.org/10.1177/1535370219867296

    • Google Scholar
87. 1. Lüders J

    (2021) Nucleating microtubules in neurons: challenges and solutions

    Developmental Neurobiology 81:273–283.

    https://doi.org/10.1002/dneu.22751

    • PubMed
    • Google Scholar
88. 1. Lüders J
    2. Stearns T

    (2007) Microtubule-organizing centres: a re-evaluation

    Nature Reviews Molecular Cell Biology 8:161–167.

    https://doi.org/10.1038/nrm2100

    • PubMed
    • Google Scholar
89. 1. Mastronarde DN
    2. McDonald KL
    3. Ding R
    4. McIntosh JR

    (1993) Interpolar spindle microtubules in PTK cells

    Journal of Cell Biology 123:1475–1489.

    https://doi.org/10.1083/jcb.123.6.1475

    • PubMed
    • Google Scholar
90. 1. Mattie FJ
    2. Stackpole MM
    3. Stone MC
    4. Clippard JR
    5. Rudnick DA
    6. Qiu Y
    7. Tao J
    8. Allender DL
    9. Parmar M
    10. Rolls MM

    (2010) Directed microtubule growth, +TIPs, and kinesin-2 are required for uniform microtubule polarity in dendrites

    Current Biology 20:2169–2177.

    https://doi.org/10.1016/j.cub.2010.11.050

    • PubMed
    • Google Scholar
91. 1. Mayr MI
    2. Hümmer S
    3. Bormann J
    4. Grüner T
    5. Adio S
    6. Woehlke G
    7. Mayer TU

    (2007) The human kinesin Kif18A is a motile microtubule depolymerase essential for chromosome congression

    Current Biology 17:488–498.

    https://doi.org/10.1016/j.cub.2007.02.036

    • PubMed
    • Google Scholar
92. 1. McNally K
    2. Audhya A
    3. Oegema K
    4. McNally FJ

    (2006) Katanin controls mitotic and meiotic spindle length

    Journal of Cell Biology 175:881–891.

    https://doi.org/10.1083/jcb.200608117

    • Google Scholar
93. 1. McNally FJ
    2. Roll-Mecak A

    (2018) Microtubule-severing enzymes: from cellular functions to molecular mechanism

    Journal of Cell Biology 217:4057–4069.

    https://doi.org/10.1083/jcb.201612104

    • Google Scholar
94. 1. Miller PM
    2. Folkmann AW
    3. Maia AR
    4. Efimova N
    5. Efimov A
    6. Kaverina I

    (2009) Golgi-derived CLASP-dependent microtubules control golgi organization and polarized trafficking in motile cells

    Nature Cell Biology 11:1069–1080.

    https://doi.org/10.1038/ncb1920

    • PubMed
    • Google Scholar
95. 1. Mitchison T
    2. Wühr M
    3. Nguyen P
    4. Ishihara K
    5. Groen A
    6. Field CM

    (2012) Growth, interaction, and positioning of microtubule asters in extremely large vertebrate embryo cells

    Cytoskeleton 69:738–750.

    https://doi.org/10.1002/cm.21050

    • PubMed
    • Google Scholar
96. 1. Mitchison T
    2. Kirschner M

    (1984) Dynamic instability of microtubule growth

    Nature 312:237–242.

    https://doi.org/10.1038/312237a0

    • PubMed
    • Google Scholar
97. 1. Molodtsov MI
    2. Mieck C
    3. Dobbelaere J
    4. Dammermann A
    5. Westermann S
    6. Vaziri A

    (2016) A Force-Induced directional switch of a molecular motor enables parallel microtubule bundle formation

    Cell 167:539–552.

    https://doi.org/10.1016/j.cell.2016.09.029

    • Google Scholar
98. 1. Murata T
    2. Sonobe S
    3. Baskin TI
    4. Hyodo S
    5. Hasezawa S
    6. Nagata T
    7. Horio T
    8. Hasebe M

    (2005) Microtubule-dependent microtubule nucleation based on recruitment of gamma-tubulin in higher plants

    Nature Cell Biology 7:961–968.

    https://doi.org/10.1038/ncb1306

    • PubMed
    • Google Scholar
99. 1. Murphy SM
    2. Preble AM
    3. Patel UK
    4. O'Connell KL
    5. Dias DP
    6. Moritz M
    7. Agard D
    8. Stults JT
    9. Stearns T

    (2001) GCP5 and GCP6: two new members of the human gamma-tubulin complex

    Molecular Biology of the Cell 12:3340–3352.

    https://doi.org/10.1091/mbc.12.11.3340

    • PubMed
    • Google Scholar
100. 1. Musa H
     2. Orton C
     3. Morrison EE
     4. Peckham M

     (2003) Microtubule assembly in cultured myoblasts and myotubes following nocodazole induced microtubule depolymerisation

     Journal of Muscle Research and Cell Motility 24:301–308.

     https://doi.org/10.1023/A:1025477807393

     • PubMed
     • Google Scholar
101. 1. Muto E
     2. Sakai H
     3. Kaseda K

     (2005) Long-range cooperative binding of kinesin to a microtubule in the presence of ATP

     Journal of Cell Biology 168:691–696.

     https://doi.org/10.1083/jcb.200409035

     • PubMed
     • Google Scholar
102. 1. Nakamura M
     2. Ehrhardt DW
     3. Hashimoto T

     (2010) Microtubule and katanin-dependent dynamics of microtubule nucleation complexes in the acentrosomal Arabidopsis cortical array

     Nature Cell Biology 12:1064–1070.

     https://doi.org/10.1038/ncb2110

     • PubMed
     • Google Scholar
103. 1. Nakamura M
     2. Lindeboom JJ
     3. Saltini M
     4. Mulder BM
     5. Ehrhardt DW

     (2018) SPR2 protects minus ends to promote severing and reorientation of plant cortical microtubule arrays

     Journal of Cell Biology 217:915–927.

     https://doi.org/10.1083/jcb.201708130

     • PubMed
     • Google Scholar
104. 1. Nakamura M
     2. Hashimoto T

     (2009) A mutation in the Arabidopsis gamma-tubulin-containing complex causes helical growth and abnormal microtubule branching

     Journal of Cell Science 122:2208–2217.

     https://doi.org/10.1242/jcs.044131

     • PubMed
     • Google Scholar
105. 1. Needleman DJ
     2. Ojeda-Lopez MA
     3. Raviv U
     4. Ewert K
     5. Jones JB
     6. Miller HP
     7. Wilson L
     8. Safinya CR

     (2004a) Synchrotron X-ray diffraction study of microtubules buckling and bundling under osmotic stress: a probe of interprotofilament interactions

     Physical Review Letters 93:198104.

     https://doi.org/10.1103/PhysRevLett.93.198104

     • PubMed
     • Google Scholar
106. 1. Needleman DJ
     2. Ojeda-Lopez MA
     3. Raviv U
     4. Miller HP
     5. Wilson L
     6. Safinya CR

     (2004b) Higher-order assembly of microtubules by counterions: from hexagonal bundles to living necklaces

     PNAS 101:16099–16103.

     https://doi.org/10.1073/pnas.0406076101

     • PubMed
     • Google Scholar
107. 1. Needleman D
     2. Dogic Z

     (2017) Active matter at the interface between materials science and cell biology

     Nature Reviews Materials 2:1–14.

     https://doi.org/10.1038/natrevmats.2017.48

     • Google Scholar
108. 1. Nixon FM
     2. Gutiérrez-Caballero C
     3. Hood FE
     4. Booth DG
     5. Prior IA
     6. Royle SJ

     (2015) The mesh is a network of microtubule connectors that stabilizes individual kinetochore fibers of the mitotic spindle

     eLife 4:e07635.

     https://doi.org/10.7554/eLife.07635

     • PubMed
     • Google Scholar
109. 1. Norris SR
     2. Jung S
     3. Singh P
     4. Strothman CE
     5. Erwin AL
     6. Ohi MD
     7. Zanic M
     8. Ohi R

     (2018) Microtubule minus-end aster organization is driven by processive HSET-tubulin clusters

     Nature communications 9:2659.

     https://doi.org/10.1038/s41467-018-04991-2

     • PubMed
     • Google Scholar
110. 1. Noujaim M
     2. Bechstedt S
     3. Wieczorek M
     4. Brouhard GJ

     (2014) Microtubules accelerate the kinase activity of Aurora-B by a reduction in dimensionality

     PLOS ONE 9:e86786.

     https://doi.org/10.1371/journal.pone.0086786

     • PubMed
     • Google Scholar
111. 1. O. WASTENEYS G
     2. E. WILLIAMSON R

     (1989) Reassembly of microtubules in Nitella tasmanica: assembly of cortical microtubules in branching clusters and its relevance to steady-state microtubule assembly

     Journal of Cell Science 93:705–714.

     https://doi.org/10.1242/jcs.93.4.705

     • Google Scholar
112. 1. Odde DJ
     2. Ma L
     3. Briggs AH
     4. DeMarco A
     5. Kirschner MW

     (1999)

     Microtubule bending and breaking in living fibroblast cells

     Journal of Cell Science 112:3283–3288.

     • PubMed
     • Google Scholar
113. 1. Oegema K
     2. Wiese C
     3. Martin OC
     4. Milligan RA
     5. Iwamatsu A
     6. Mitchison TJ
     7. Zheng Y

     (1999) Characterization of two related Drosophila gamma-tubulin complexes that differ in their ability to nucleate microtubules

     The Journal of Cell Biology 144:721–733.

     https://doi.org/10.1083/jcb.144.4.721

     • PubMed
     • Google Scholar
114. 1. Oh D
     2. Yu CH
     3. Needleman DJ

     (2016) Spatial organization of the Ran pathway by microtubules in mitosis

     PNAS 113:8729–8734.

     https://doi.org/10.1073/pnas.1607498113

     • PubMed
     • Google Scholar
115. 1. Pamula MC
     2. Carlini L
     3. Forth S
     4. Verma P
     5. Suresh S
     6. Legant WR
     7. Khodjakov A
     8. Betzig E
     9. Kapoor TM

     (2019) High-resolution imaging reveals how the spindle midzone impacts chromosome movement

     Journal of Cell Biology 218:2529–2544.

     https://doi.org/10.1083/jcb.201904169

     • Google Scholar
116. 1. Paredez AR
     2. Somerville CR
     3. Ehrhardt DW

     (2006) Visualization of cellulose synthase demonstrates functional association with microtubules

     Science 312:1491–1495.

     https://doi.org/10.1126/science.1126551

     • PubMed
     • Google Scholar
117. 1. Paz J
     2. Lüders J

     (2018) Microtubule-Organizing Centers: Towards a Minimal Parts List

     Trends in Cell Biology 28:176–187.

     https://doi.org/10.1016/j.tcb.2017.10.005

     • PubMed
     • Google Scholar
118. 1. Peet DR
     2. Burroughs NJ
     3. Cross RA

     (2018) Kinesin expands and stabilizes the GDP-microtubule lattice

     Nature Nanotechnology 13:386–391.

     https://doi.org/10.1038/s41565-018-0084-4

     • PubMed
     • Google Scholar
119. 1. Peterman EJ
     2. Scholey JM

     (2009) Mitotic microtubule crosslinkers: insights from mechanistic studies

     Current Biology: CB 19:R1089–R1094.

     https://doi.org/10.1016/j.cub.2009.10.047

     • PubMed
     • Google Scholar
120. 1. Petry S
     2. Groen AC
     3. Ishihara K
     4. Mitchison TJ
     5. Vale RD

     (2013) Branching microtubule nucleation in Xenopus egg extracts mediated by augmin and TPX2

     Cell 152:768–777.

     https://doi.org/10.1016/j.cell.2012.12.044

     • PubMed
     • Google Scholar
121. 1. Petry S
     2. Vale RD

     (2015) Microtubule nucleation at the centrosome and beyond

     Nature Cell Biology 17:1089–1093.

     https://doi.org/10.1038/ncb3220

     • PubMed
     • Google Scholar
122. 1. Pollard TD
     2. O'Shaughnessy B

     (2019) Molecular Mechanism of Cytokinesis

     Annual Review of Biochemistry 88:661–689.

     https://doi.org/10.1146/annurev-biochem-062917-012530

     • PubMed
     • Google Scholar
123. 1. Portran D
     2. Schaedel L
     3. Xu Z
     4. Théry M
     5. Nachury MV

     (2017) Tubulin acetylation protects long-lived microtubules against mechanical ageing

     Nature Cell Biology 19:391–398.

     https://doi.org/10.1038/ncb3481

     • PubMed
     • Google Scholar
124. 1. Pringle J
     2. Muthukumar A
     3. Tan A
     4. Crankshaw L
     5. Conway L
     6. Ross JL

     (2013) Microtubule organization by kinesin motors and microtubule crosslinking protein MAP65

     Journal of Physics: Condensed Matter 25:374103.

     https://doi.org/10.1088/0953-8984/25/37/374103

     • Google Scholar
125. 1. Rai A
     2. Liu T
     3. Glauser S
     4. Katrukha EA
     5. Estévez-Gallego J
     6. Rodríguez-García R
     7. Fang WS
     8. Díaz JF
     9. Steinmetz MO
     10. Altmann KH
     11. Kapitein LC
     12. Moores CA
     13. Akhmanova A

     (2020) Taxanes convert regions of perturbed microtubule growth into rescue sites

     Nature Materials 19:355–365.

     https://doi.org/10.1038/s41563-019-0546-6

     • PubMed
     • Google Scholar
126. Preprint

     1. Rai A
     2. Liu T
     3. Katrukha EA
     4. Estévez-Gallego J
     5. Paterson I
     6. Diaz F
     7. Kapitein LC
     8. Moores CA
     9. Akhmanova A

     (2021) Microtubule lattice defects promote catastrophes

     bioRxiv.

     https://doi.org/10.1101/2021.02.11.430743

     • Google Scholar
127. 1. Rickman J
     2. Nédélec F
     3. Surrey T

     (2019) Effects of spatial dimensionality and steric interactions on microtubule-motor self-organization

     Physical Biology 16:046004.

     https://doi.org/10.1088/1478-3975/ab0fb1

     • PubMed
     • Google Scholar
128. 1. Rizk RS
     2. Discipio KA
     3. Proudfoot KG
     4. Gupta ML

     (2014) The kinesin-8 Kip3 scales anaphase spindle length by suppression of midzone microtubule polymerization

     The Journal of Cell Biology 204:965–975.

     https://doi.org/10.1083/jcb.201312039

     • PubMed
     • Google Scholar
129. 1. Robison P
     2. Caporizzo MA
     3. Ahmadzadeh H
     4. Bogush AI
     5. Chen CY
     6. Margulies KB
     7. Shenoy VB
     8. Prosser BL

     (2016) Detyrosinated microtubules buckle and bear load in contracting cardiomyocytes

     Science 352:aaf0659.

     https://doi.org/10.1126/science.aaf0659

     • PubMed
     • Google Scholar
130. 1. Roll-Mecak A
     2. Vale RD

     (2006) Making more microtubules by severing: a common theme of noncentrosomal microtubule arrays?

     The Journal of Cell Biology 175:849–851.

     https://doi.org/10.1083/jcb.200611149

     • PubMed
     • Google Scholar
131. 1. Roostalu J
     2. Hentrich C
     3. Bieling P
     4. Telley IA
     5. Schiebel E
     6. Surrey T

     (2011) Directional switching of the kinesin Cin8 through motor coupling

     Science 332:94–99.

     https://doi.org/10.1126/science.1199945

     • PubMed
     • Google Scholar
132. 1. Roostalu J
     2. Rickman J
     3. Thomas C
     4. Nédélec F
     5. Surrey T

     (2018) Determinants of Polar versus Nematic Organization in Networks of Dynamic Microtubules and Mitotic Motors

     Cell 175:e14.

     https://doi.org/10.1016/j.cell.2018.09.029

     • PubMed
     • Google Scholar
133. 1. Roostalu J
     2. Surrey T

     (2017) Microtubule nucleation: beyond the template

     Nature Reviews. Molecular Cell Biology 18:702–710.

     https://doi.org/10.1038/nrm.2017.75

     • PubMed
     • Google Scholar
134. 1. Sánchez I
     2. Dynlacht BD

     (2016) Cilium assembly and disassembly

     Nature Cell Biology 18:711–717.

     https://doi.org/10.1038/ncb3370

     • PubMed
     • Google Scholar
135. 1. Sánchez-Huertas C
     2. Freixo F
     3. Viais R
     4. Lacasa C
     5. Soriano E
     6. Lüders J

     (2016) Non-centrosomal nucleation mediated by augmin organizes microtubules in post-mitotic neurons and controls axonal microtubule polarity

     Nature Communications 7:12187.

     https://doi.org/10.1038/ncomms12187

     • PubMed
     • Google Scholar
136. 1. Schaedel L
     2. John K
     3. Gaillard J
     4. Nachury MV
     5. Blanchoin L
     6. Théry M

     (2015) Microtubules self-repair in response to mechanical stress

     Nature Materials 14:1156–1163.

     https://doi.org/10.1038/nmat4396

     • PubMed
     • Google Scholar
137. 1. Schaedel L
     2. Triclin S
     3. Chrétien D
     4. Abrieu A
     5. Aumeier C
     6. Gaillard J
     7. Blanchoin L
     8. Théry M
     9. John K

     (2019) Lattice defects induce microtubule self-renewal

     Nature Physics 15:830–838.

     https://doi.org/10.1038/s41567-019-0542-4

     • PubMed
     • Google Scholar
138. 1. Schmidt-Cernohorska M
     2. Zhernov I
     3. Steib E
     4. Le Guennec M
     5. Achek R
     6. Borgers S
     7. Demurtas D
     8. Mouawad L
     9. Lansky Z
     10. Hamel V
     11. Guichard P

     (2019) Flagellar microtubule doublet assembly in vitro reveals a regulatory role of tubulin C-terminal tails

     Science 363:285–288.

     https://doi.org/10.1126/science.aav2567

     • PubMed
     • Google Scholar
139. 1. Scholey J
     2. Civelekoglu-Scholey G
     3. Brust-Mascher I

     (2016) Anaphase B

     Biology 5:51.

     https://doi.org/10.3390/biology5040051

     • Google Scholar
140. 1. Schwarz PM
     2. Liggins JR
     3. Ludueña RF

     (1998) Beta-tubulin isotypes purified from bovine brain have different relative stabilities

     Biochemistry 37:4687–4692.

     https://doi.org/10.1021/bi972763d

     • PubMed
     • Google Scholar
141. 1. Seetharaman S
     2. Etienne-Manneville S

     (2020) Cytoskeletal Crosstalk in Cell Migration

     Trends in Cell Biology 30:720–735.

     https://doi.org/10.1016/j.tcb.2020.06.004

     • PubMed
     • Google Scholar
142. 1. Serrano L
     2. de la Torre J
     3. Maccioni RB
     4. Avila J

     (1984) Involvement of the carboxyl-terminal domain of tubulin in the regulation of its assembly

     PNAS 81:5989–5993.

     https://doi.org/10.1073/pnas.81.19.5989

     • PubMed
     • Google Scholar
143. 1. She ZY
     2. Wei YL
     3. Lin Y
     4. Li YL
     5. Lu MH

     (2019) Mechanisms of the Ase1/PRC1/MAP65 family in central spindle assembly

     Biological Reviews of the Cambridge Philosophical Society 94:2033–2048.

     https://doi.org/10.1111/brv.12547

     • PubMed
     • Google Scholar
144. 1. Shima T
     2. Morikawa M
     3. Kaneshiro J
     4. Kambara T
     5. Kamimura S
     6. Yagi T
     7. Iwamoto H
     8. Uemura S
     9. Shigematsu H
     10. Shirouzu M
     11. Ichimura T
     12. Watanabe TM
     13. Nitta R
     14. Okada Y
     15. Hirokawa N

     (2018) Kinesin-binding-triggered conformation switching of microtubules contributes to polarized transport

     The Journal of Cell Biology 217:4164–4183.

     https://doi.org/10.1083/jcb.201711178

     • PubMed
     • Google Scholar
145. 1. Shimamoto Y
     2. Forth S
     3. Kapoor TM

     (2015) Measuring Pushing and Braking Forces Generated by Ensembles of Kinesin-5 Crosslinking Two Microtubules

     Developmental Cell 34:669–681.

     https://doi.org/10.1016/j.devcel.2015.08.017

     • PubMed
     • Google Scholar
146. 1. Sirajuddin M
     2. Rice LM
     3. Vale RD

     (2014) Regulation of microtubule motors by tubulin isotypes and post-translational modifications

     Nature Cell Biology 16:335–344.

     https://doi.org/10.1038/ncb2920

     • PubMed
     • Google Scholar
147. 1. Song JG
     2. King MR
     3. Zhang R
     4. Kadzik RS
     5. Thawani A
     6. Petry S

     (2018) Mechanism of how augmin directly targets the γ-tubulin ring complex to microtubules

     The Journal of Cell Biology 217:2417–2428.

     https://doi.org/10.1083/jcb.201711090

     • PubMed
     • Google Scholar
148. 1. Song Y
     2. Brady ST

     (2015) Post-translational modifications of tubulin: pathways to functional diversity of microtubules

     Trends in Cell Biology 25:125–136.

     https://doi.org/10.1016/j.tcb.2014.10.004

     • PubMed
     • Google Scholar
149. 1. Srayko M
     2. O'toole ET
     3. Hyman AA
     4. Müller-Reichert T

     (2006) Katanin disrupts the microtubule lattice and increases polymer number in C. elegans meiosis

     Current Biology: CB 16:1944–1949.

     https://doi.org/10.1016/j.cub.2006.08.029

     • PubMed
     • Google Scholar
150. 1. Stairs CW
     2. Ettema TJG

     (2020) The Archaeal Roots of the Eukaryotic Dynamic Actin Cytoskeleton

     Current Biology: CB 30:R521–R526.

     https://doi.org/10.1016/j.cub.2020.02.074

     • PubMed
     • Google Scholar
151. 1. Stanhope KT
     2. Yadav V
     3. Santangelo CD
     4. Ross JL

     (2017) Contractility in an extensile system

     Soft Matter 13:4268–4277.

     https://doi.org/10.1039/c7sm00449d

     • PubMed
     • Google Scholar
152. 1. Stone MC
     2. Roegiers F
     3. Rolls MM

     (2008) Microtubules have opposite orientation in axons and dendrites of Drosophila neurons

     Molecular Biology of the Cell 19:4122–4129.

     https://doi.org/10.1091/mbc.e07-10-1079

     • PubMed
     • Google Scholar
153. 1. Subramanian R
     2. Ti SC
     3. Tan L
     4. Darst SA
     5. Kapoor TM

     (2013) Marking and measuring single microtubules by PRC1 and kinesin-4

     Cell 154:377–390.

     https://doi.org/10.1016/j.cell.2013.06.021

     • PubMed
     • Google Scholar
154. 1. Subramanian R
     2. Kapoor TM

     (2012) Building complexity: insights into self-organized assembly of microtubule-based architectures

     Developmental Cell 23:874–885.

     https://doi.org/10.1016/j.devcel.2012.10.011

     • PubMed
     • Google Scholar
155. 1. Szyk A
     2. Deaconescu AM
     3. Piszczek G
     4. Roll-Mecak A

     (2011) Tubulin tyrosine ligase structure reveals adaptation of an ancient fold to bind and modify tubulin

     Nature Structural & Molecular Biology 18:1250–1258.

     https://doi.org/10.1038/nsmb.2148

     • PubMed
     • Google Scholar
156. 1. Szyk A
     2. Deaconescu AM
     3. Spector J
     4. Goodman B
     5. Valenstein ML
     6. Ziolkowska NE
     7. Kormendi V
     8. Grigorieff N
     9. Roll-Mecak A

     (2014) Molecular basis for age-dependent microtubule acetylation by tubulin acetyltransferase

     Cell 157:1405–1415.

     https://doi.org/10.1016/j.cell.2014.03.061

     • PubMed
     • Google Scholar
157. 1. Tan R
     2. Foster PJ
     3. Needleman DJ
     4. McKenney RJ

     (2018) Cooperative Accumulation of Dynein-Dynactin at Microtubule Minus-Ends Drives Microtubule Network Reorganization

     Developmental Cell 44:233–247.

     https://doi.org/10.1016/j.devcel.2017.12.023

     • PubMed
     • Google Scholar
158. 1. Tariq A
     2. Green L
     3. Jeynes JCG
     4. Soeller C
     5. Wakefield JG

     (2020) In vitro reconstitution of branching microtubule nucleation

     eLife 9:e49769.

     https://doi.org/10.7554/eLife.49769

     • PubMed
     • Google Scholar
159. 1. Tian J
     2. Kong Z

     (2019) The role of the augmin complex in establishing microtubule arrays

     Journal of Experimental Botany 70:3035–3041.

     https://doi.org/10.1093/jxb/erz123

     • PubMed
     • Google Scholar
160. 1. Tropini C
     2. Roth EA
     3. Zanic M
     4. Gardner MK
     5. Howard J

     (2012) Islands containing slowly hydrolyzable GTP analogs promote microtubule rescues

     PLOS ONE 7:e30103.

     https://doi.org/10.1371/journal.pone.0030103

     • PubMed
     • Google Scholar
161. 1. Uehara R
     2. Kamasaki T
     3. Hiruma S
     4. Poser I
     5. Yoda K
     6. Yajima J
     7. Gerlich DW
     8. Goshima G

     (2016) Augmin shapes the anaphase spindle for efficient cytokinetic furrow ingression and abscission

     Molecular Biology of the Cell 27:812–827.

     https://doi.org/10.1091/mbc.E15-02-0101

     • PubMed
     • Google Scholar
162. 1. Uehara R
     2. Goshima G

     (2010) Functional central spindle assembly requires de novo microtubule generation in the interchromosomal region during anaphase

     The Journal of Cell Biology 191:259–267.

     https://doi.org/10.1083/jcb.201004150

     • PubMed
     • Google Scholar
163. 1. Valenstein ML
     2. Roll-Mecak A

     (2016) Graded Control of Microtubule Severing by Tubulin Glutamylation

     Cell 164:911–921.

     https://doi.org/10.1016/j.cell.2016.01.019

     • PubMed
     • Google Scholar
164. 1. van Beuningen SF
     2. Hoogenraad CC

     (2016) Neuronal polarity: remodeling microtubule organization

     Current Opinion in Neurobiology 39:1–7.

     https://doi.org/10.1016/j.conb.2016.02.003

     • PubMed
     • Google Scholar
165. 1. van de Willige D
     2. Hoogenraad CC
     3. Akhmanova A

     (2016) Microtubule plus-end tracking proteins in neuronal development

     Cellular and Molecular Life Sciences 73:2053–2077.

     https://doi.org/10.1007/s00018-016-2168-3

     • PubMed
     • Google Scholar
166. 1. Varga V
     2. Helenius J
     3. Tanaka K
     4. Hyman AA
     5. Tanaka TU
     6. Howard J

     (2006) Yeast kinesin-8 depolymerizes microtubules in a length-dependent manner

     Nature Cell Biology 8:957–962.

     https://doi.org/10.1038/ncb1462

     • PubMed
     • Google Scholar
167. 1. Varga V
     2. Leduc C
     3. Bormuth V
     4. Diez S
     5. Howard J

     (2009) Kinesin-8 motors act cooperatively to mediate length-dependent microtubule depolymerization

     Cell 138:1174–1183.

     https://doi.org/10.1016/j.cell.2009.07.032

     • PubMed
     • Google Scholar
168. 1. Vemu A
     2. Szczesna E
     3. Zehr EA
     4. Spector JO
     5. Grigorieff N
     6. Deaconescu AM
     7. Roll-Mecak A

     (2018) Severing enzymes amplify microtubule arrays through lattice GTP-tubulin incorporation

     Science 361:eaau1504.

     https://doi.org/10.1126/science.aau1504

     • PubMed
     • Google Scholar
169. 1. Verhey KJ
     2. Gaertig J

     (2007) The tubulin code

     Cell Cycle 6:2152–2160.

     https://doi.org/10.4161/cc.6.17.4633

     • PubMed
     • Google Scholar
170. 1. Vilfan A
     2. Frey E
     3. Schwabl F
     4. Thormählen M
     5. Song YH
     6. Mandelkow E

     (2001) Dynamics and cooperativity of microtubule decoration by the motor protein kinesin

     Journal of Molecular Biology 312:1011–1026.

     https://doi.org/10.1006/jmbi.2001.5020

     • PubMed
     • Google Scholar
171. 1. Vukušić K
     2. Buđa R
     3. Tolić IM

     (2019) Force-generating mechanisms of anaphase in human cells

     Journal of Cell Science 132:jcs231985.

     https://doi.org/10.1242/jcs.231985

     • PubMed
     • Google Scholar
172. 1. Walker RA
     2. O'Brien ET
     3. Pryer NK
     4. Soboeiro MF
     5. Voter WA
     6. Erickson HP
     7. Salmon ED

     (1988) Dynamic instability of individual microtubules analyzed by video light microscopy: rate constants and transition frequencies

     The Journal of Cell Biology 107:1437–1448.

     https://doi.org/10.1083/jcb.107.4.1437

     • PubMed
     • Google Scholar
173. 1. Wang C
     2. Liu W
     3. Wang G
     4. Li J
     5. Dong L
     6. Han L
     7. Wang Q
     8. Tian J
     9. Yu Y
     10. Gao C
     11. Kong Z

     (2017) KTN80 confers precision to microtubule severing by specific targeting of katanin complexes in plant cells

     The EMBO Journal 36:3435–3447.

     https://doi.org/10.15252/embj.201796823

     • PubMed
     • Google Scholar
174. 1. Wasteneys GO
     2. Ambrose JC

     (2009) Spatial organization of plant cortical microtubules: close encounters of the 2D kind

     Trends in Cell Biology 19:62–71.

     https://doi.org/10.1016/j.tcb.2008.11.004

     • PubMed
     • Google Scholar
175. 1. Watanabe T
     2. Noritake J
     3. Kaibuchi K

     (2005) Regulation of microtubules in cell migration

     Trends in Cell Biology 15:76–83.

     https://doi.org/10.1016/j.tcb.2004.12.006

     • PubMed
     • Google Scholar
176. 1. Waterman-Storer CM
     2. Salmon ED

     (1997) Actomyosin-based retrograde flow of microtubules in the lamella of migrating epithelial cells influences microtubule dynamic instability and turnover and is associated with microtubule breakage and treadmilling

     The Journal of Cell Biology 139:417–434.

     https://doi.org/10.1083/jcb.139.2.417

     • PubMed
     • Google Scholar
177. 1. Wieczorek M
     2. Urnavicius L
     3. Ti S-C
     4. Molloy KR
     5. Chait BT
     6. Kapoor TM

     (2020) Asymmetric molecular architecture of the human γ-Tubulin ring complex

     Cell 180:165–175.

     https://doi.org/10.1016/j.cell.2019.12.007

     • Google Scholar
178. 1. Wightman R
     2. Turner SR

     (2007) Severing at sites of microtubule crossover contributes to microtubule alignment in cortical arrays

     The Plant Journal 52:742–751.

     https://doi.org/10.1111/j.1365-313X.2007.03271.x

     • PubMed
     • Google Scholar
179. Preprint

     1. Wijeratne SS
     2. Fiorenza SA
     3. Subramanian R
     4. Betterton MD

     (2020) Motor guidance by long-range communication through the microtubule highway

     bioRxiv.

     https://doi.org/10.1101/2020.12.23.424221

     • Google Scholar
180. 1. Wijeratne S
     2. Subramanian R

     (2018) Geometry of antiparallel microtubule bundles regulates relative sliding and stalling by PRC1 and Kif4A

     eLife 7:e32595.

     https://doi.org/10.7554/eLife.32595

     • PubMed
     • Google Scholar
181. 1. Wloga D
     2. Joachimiak E
     3. Louka P
     4. Gaertig J

     (2017) Posttranslational modifications of tubulin and cilia

     Cold Spring Harbor Perspectives in Biology 9:a028159.

     https://doi.org/10.1101/cshperspect.a028159

     • PubMed
     • Google Scholar
182. 1. Wu J
     2. Akhmanova A

     (2017) Microtubule-Organizing centers

     Annual Review of Cell and Developmental Biology 33:51–75.

     https://doi.org/10.1146/annurev-cellbio-100616-060615

     • PubMed
     • Google Scholar
183. 1. Yagi N
     2. Matsunaga S
     3. Hashimoto T

     (2018) Insights into cortical microtubule nucleation and dynamics in Arabidopsis leaf cells

     Journal of Cell Science 131:jcs203778.

     https://doi.org/10.1242/jcs.203778

     • PubMed
     • Google Scholar
184. 1. Yi P
     2. Goshima G

     (2018) Microtubule nucleation and organization without centrosomes

     Current Opinion in Plant Biology 46:1–7.

     https://doi.org/10.1016/j.pbi.2018.06.004

     • PubMed
     • Google Scholar
185. 1. Zandomeni K
     2. Schopfer P

     (1993) Reorientation of microtubules at the outer epidermal wall of maize coleoptiles by Phytochrome, blue-light photoreceptor, and auxin

     Protoplasma 173:103–112.

     https://doi.org/10.1007/BF01378999

     • Google Scholar
186. 1. Zanic M
     2. Widlund PO
     3. Hyman AA
     4. Howard J

     (2013) Synergy between XMAP215 and EB1 increases microtubule growth rates to physiological levels

     Nature Cell Biology 15:688–693.

     https://doi.org/10.1038/ncb2744

     • PubMed
     • Google Scholar
187. 1. Zhang Q
     2. Fishel E
     3. Bertroche T
     4. Dixit R

     (2013) Microtubule severing at crossover sites by katanin generates ordered cortical microtubule arrays in Arabidopsis

     Current Biology: CB 23:2191–2195.

     https://doi.org/10.1016/j.cub.2013.09.018

     • PubMed
     • Google Scholar
188. 1. Zheng Y
     2. Wong ML
     3. Alberts B
     4. Mitchison T

     (1995) Nucleation of microtubule assembly by a gamma-tubulin-containing ring complex

     Nature 378:578–583.

     https://doi.org/10.1038/378578a0

     • PubMed
     • Google Scholar

Author details

1. Nandini Mani

   1. Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Department of Genetics, Harvard Medical School, Boston, United States

   Contribution

   Conceptualization, Visualization, Writing - original draft, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-2025-9103

2. Sithara S Wijeratne

   1. Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Department of Genetics, Harvard Medical School, Boston, United States

   Contribution

   Visualization, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3935-926X

3. Radhika Subramanian

   1. Department of Molecular Biology, Massachusetts General Hospital, Boston, United States
   2. Department of Genetics, Harvard Medical School, Boston, United States

   Contribution

   Conceptualization, Funding acquisition, Writing - original draft, Writing - review and editing

   For correspondence

   radhika@molbio.mgh.harvard.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3011-9403

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