Abstract

Microtubules are dynamic polymers that grow and shrink through addition or loss of tubulin subunits at their ends. Microtubule ends generate mechanical force that moves chromosomes and cellular organelles, and provides mechanical tension. Recent literature describes a number of proteins and protein complexes that couple dynamics of microtubule ends to movements of their cellular cargoes. These ‘couplers’ are quite diverse in their microtubule-binding domains (MTBDs), while sharing similarity in function, but a systematic understanding of the principles underlying their activity is missing. Here, I review various types of microtubule couplers, focusing on their essential activities: ability to follow microtubule ends and capture microtubule-generated force. Most of the couplers require presence of unstructured positively charged sequences and multivalency in their microtubule-binding sites to efficiently convert the microtubule-generated force into useful connection to a cargo. An overview of the microtubule features supporting end-tracking and force-coupling, and the experimental methods to assess force-coupling properties is also provided.

Introduction

Cells actively rearrange themselves during the life cycle: vesicles and organelles get transported, membranes get remodelled, chromosomes get separated. Many of these movements are driven by microtubule dynamics, with cell division being the best studied context for microtubule-generated force (reviewed in [1]). Microtubules are 25-nm wide hollow tubes built from α- and β-tubulin heterodimers, and are constantly growing or shortening from their ends through addition or removal of tubulin dimers, respectively. Microtubule dynamics are linked to the nucleotide hydrolysis: GTP-bound tubulin binds to the end of the microtubule and later hydrolyzes GTP to GDP inside the microtubule lattice. If GDP-containing tubulin stochastically crowds out the GTP-tubulin, the microtubule end switches to shortening (reviewed in [2]).

Tubulin dimers and bigger oligomers are bent in solution, but in the microtubule lattice they straighten as they bind to the neighbouring tubulin dimers [3] (Figure 1). Interestingly, the bent shapes of tubulin protofilaments at both growing and shortening microtubules are similar [3] (Figure 1). The microtubule end presents a unique interface in which transitions of tubulin shape from bent (at the end) to straight (in the lattice) and back are coupled to the energy released during GTP hydrolysis. The mechano-chemical link between GTP hydrolysis and dynamics of tubulin protofilaments allows the microtubules to exert force on a cargo with their ends (Figure 1).

Structure of a dynamic microtubule end

Figure 1
Structure of a dynamic microtubule end

(A) A microtubule end is characterized by protofilaments, linear strands of tubulin dimers, bending outward. Shortening microtubule end generates force thanks to the power strokes of individual protofilaments pushing a cargo in the direction of microtubule shortening (red arrows). Microtubule growth can also generate force (black arrow). The microtubule image is produced from coordinates obtained in a coarse-grained molecular dynamics simulation (courtesy of Nikita Gudimchuk [3]), rendered using tubulin structure from [4]. (B,C) Shapes of tubulin protofilaments obtained from cryo-electron tomographic reconstructions. Whether the microtubules are growing (B) or shrinking (C), the shapes of the tubulin curls at the microtubule ends are not significantly different (reproduced with permission from [3] ©Rockefeller University Press).

Figure 1
Structure of a dynamic microtubule end

(A) A microtubule end is characterized by protofilaments, linear strands of tubulin dimers, bending outward. Shortening microtubule end generates force thanks to the power strokes of individual protofilaments pushing a cargo in the direction of microtubule shortening (red arrows). Microtubule growth can also generate force (black arrow). The microtubule image is produced from coordinates obtained in a coarse-grained molecular dynamics simulation (courtesy of Nikita Gudimchuk [3]), rendered using tubulin structure from [4]. (B,C) Shapes of tubulin protofilaments obtained from cryo-electron tomographic reconstructions. Whether the microtubules are growing (B) or shrinking (C), the shapes of the tubulin curls at the microtubule ends are not significantly different (reproduced with permission from [3] ©Rockefeller University Press).

Shortening microtubules provide the driving force for chromosome segregation in mitosis [1,5]. Microtubule growth can also generate force (reviewed in [6]) and redistribute membranous networks, such as ER [7–9] or mitochondria [10,11]. Microtubule-generated force is transmitted to the cellular cargoes though microtubule couplers: proteins and protein complexes that harness the energy released during stochastic cycles of microtubule growth and shortening, and convert it into organized motility of cellular cargoes.

In this review, I focus on the ability of microtubule couplers to keep attachment to the microtubule end as it shortens, an activity I will refer to as ‘end-tracking’. Persistent, or processive, end-tracking requires that the coupler constantly breaks connections with a protofilament or a microtubule and rebinds again more distal from the initial position of the microtubule end, so multivalency of microtubule-binding sites is important. For many of the couplers described below, the multivalency is provided by the presence of multiple copies of a microtubule-binding protein bound to a cargo. Since the first demonstrations of the end-tracking by purified chromosomes [12], only a few recombinant proteins or protein complexes were shown to support motility with the shortening microtubule ends (summarized in Figure 2). These couplers have a diverse set of microtubule-binding domains (MTBDs), including globular domains with specific folds, completely unstructured microtubule-binding sequences, or combinations of folded and unstructured domains. Two characteristics that seem common to most of microtubule couplers are (1) disordered positively charged sequences (summarized in Table 1), and (2) multiple MTBDs to follow microtubule shortening.

Overview of microtubule end-couplers

Figure 2
Overview of microtubule end-couplers

(A) A summary of end-tracking and force-coupling complexes discussed in the text. Graphs show the probability of disordered sequence from 0 (improbable) to 1 (probable) on y-axis and sequence position on x-axis, as predicted by DISOPRED3 [13]. Red colour denotes the known microtubule-binding regions in the cartoons and on top of the disorder prediction plots. See text for detailed discussion of the differences between the rupture and stalling forces. ‘n.d.’ equals ‘not determined’.

Figure 2
Overview of microtubule end-couplers

(A) A summary of end-tracking and force-coupling complexes discussed in the text. Graphs show the probability of disordered sequence from 0 (improbable) to 1 (probable) on y-axis and sequence position on x-axis, as predicted by DISOPRED3 [13]. Red colour denotes the known microtubule-binding regions in the cartoons and on top of the disorder prediction plots. See text for detailed discussion of the differences between the rupture and stalling forces. ‘n.d.’ equals ‘not determined’.

Table 1
Length, charge, and amino acid composition of disordered microtubule-binding sequences.
Length, a.a.pIPositive a.a. (RKH, %)Polar a.a. (GQNS, %)Hydrophobic a.a. (AVIL, %)Negative a.a. (DE, %)
Ndc80 human tail 81 10.84 21.0 39.5 16.0 6.2 
Ndc80 yeast tail 80 11.88 17.5 36.3 17.5 3.8 
CENP-Q tail 67 10.43 31.3 29.9 20.9 9.0 
Dam1 C-terminus 189 10.11 20.1 32.3 20.1 11.1 
Duo1 C-terminus 68 12.61 23.5 30.9 19.1 0.0 
CENP-F C-terminus 221 10.28 17.2 29.0 22.2 9.5 
KKT4 linker 258 10.28 12.8 33.3 22.1 5.8 
MACF2 C-terminus 43 11.85 27.9 23.3 14.0 4.7 
KA7 14 10.78 50.0 0.0 50.0 0.0 
XMAP215 linker 82 9.94 17.1 31.7 14.6 7.3 
Clasp2 linker 330 10.54 15.2 38.2 25.2 7.9 
CENP-E tail 190 9.2 18.4 30.5 19.5 11.1 
Abl2 linker 131 4.84 9.9 19.1 26.0 13.0 
Length, a.a.pIPositive a.a. (RKH, %)Polar a.a. (GQNS, %)Hydrophobic a.a. (AVIL, %)Negative a.a. (DE, %)
Ndc80 human tail 81 10.84 21.0 39.5 16.0 6.2 
Ndc80 yeast tail 80 11.88 17.5 36.3 17.5 3.8 
CENP-Q tail 67 10.43 31.3 29.9 20.9 9.0 
Dam1 C-terminus 189 10.11 20.1 32.3 20.1 11.1 
Duo1 C-terminus 68 12.61 23.5 30.9 19.1 0.0 
CENP-F C-terminus 221 10.28 17.2 29.0 22.2 9.5 
KKT4 linker 258 10.28 12.8 33.3 22.1 5.8 
MACF2 C-terminus 43 11.85 27.9 23.3 14.0 4.7 
KA7 14 10.78 50.0 0.0 50.0 0.0 
XMAP215 linker 82 9.94 17.1 31.7 14.6 7.3 
Clasp2 linker 330 10.54 15.2 38.2 25.2 7.9 
CENP-E tail 190 9.2 18.4 30.5 19.5 11.1 
Abl2 linker 131 4.84 9.9 19.1 26.0 13.0 

Abbreviations: CENP, centromere protein; MACF2, microtubule-actin cross-linking factor 2.

Overview of microtubule couplers

Couplers with folded domains

Ska complex

Any generic microtubule-associated protein, including the ones without end-tracking properties, might occasionally produce motion of a cargo [1]. However, processive tracking of depolymerizing microtubule ends is rare among microtubule-associated proteins with folded MTBDs. To date the only efficient end-tracker that does not require contributions from intrinsically disordered regions is the Ska complex, a metazoan kinetochore component that is loaded onto the microtubule-kinetochore interface late in mitosis and is required for proper segregation of chromosomes [14,15]. SKA123 complex binds microtubules through the C-terminal winged-helix domain of SKA1 [16,17]. SKA3 subunit carries an extensive disordered region, which is however not required for interaction between Ska complex and microtubules [18,19]. The end-tracking ability of Ska complex is attributed to its higher affinity for bent tubulin, which is present at the microtubule ends, compared with straight tubulin in the microtubule lattice [20]. Importantly, affinity for microtubule ends alone is insufficient for end-tracking since the end-bound coupler needs to rebind as the microtubule shortens [21]. Consistently with this, Ska was shown to end-track if oligomerized on the surface of the bead [22]. In the absence of a cargo, fluorescent Ska particles were following both microtubule growth and shortening, however it is unknown if individual molecules of Ska also have this ability [20,23,24]. Enrichment of Ska at the ends of microtubules depended on positively charged amino acids in SKA1 MTBD: K223A, R155A and R236A substitutions abolished enrichment of Ska at the both growing and shortening microtubule ends, and K183/184A double substitution preferentially impaired enrichment at growing microtubule ends, while still supporting end-tracking with microtubule shortening [23]. These observations suggest that both affinity to microtubule ends, and potentially presence of multiple copies of Ska are required for efficient end-tracking.

Couplers with only unstructured microtubule-binding sequences

Dam1/DASH complex

One of the first complexes shown to follow microtubule ends autonomously was Dam1/DASH [25]. Dam1 complex is a kinetochore component present in fungi and is essential for viability of Saccharomyces cerevisiae [26]. Dam1 oligomerizes into rings around microtubules [27], but can also end-track as smaller, non-ring oligomers [28]. Dam1 and Ska complexes are considered to be functional paralogs, because they are not related and they do not coexist in the same species, while performing the same function of enhancing Ndc80-microtubule attachment in the kinetochore [22,29]. The principal unit of Dam1 is a heterodecamer which binds to microtubules through the protrusion connecting the ring with the microtubule surface [30]. This protrusion is thought to be formed by positively charged disordered C-terminal tails of Dam1 and Duo1 subunits [31–33] (Figure 2 and Table 1), and is crucial for tight binding to microtubule and efficient force-coupling [34], although the contributions from other subunits are also possible [35].

Microtubule-actin cross-linking factor 2 and (KA7)4

Two completely disordered sequences with a net positive charge have been shown to promote microtubule end-tracking and capture microtubule-generated force. One is a peptide containing 43 C-terminal amino acids of human microtubule-actin cross-linking factor 2 (MACF2/dystonin) [36]. MACF2 C-terminal peptide was present in multiple copies on the surface of a quantum dot or a glass bead enabling these cargoes to follow microtubule shortening [9]. KA74 is a tetravalent particle containing fully synthetic peptide with seven repeats of lysine and alanine [37]. All the three unstructured couplers have been also shown to capture considerable force from shortening microtubule ends [9,37], with Dam1 being the absolute champion thanks to its ring shape, also providing multivalency, and tight binding to the microtubule [38,39] (Figure 2).

Couplers with combinations of folded and unstructured domains

Ndc80 complex

Ndc80 complex is a kinetochore component in majority of eukaryotes and is essential for their viability since it provides the main link between kinetochores and microtubule ends [40,41]. Ndc80’s MTBD consists of a folded calponin homology (CH) domain and an unstructured N-terminal tail, both containing positively charged amino acids (Table 1). Ndc80 tail also contains multiple sites for phosphorylation by Aurora B kinase [42,43]; Ndc80 with phosphorylated tail binds microtubules with lower affinity, highlighting the importance of the charge in Ndc80 tail for the kinetochore function [44]. The contribution of multivalency to Ndc80’s end-tracking has been appreciated for a decade [45,46], but it was only recently established that as few as two Ndc80 copies joined together are sufficient to end-track, while multiple Ndc80 di- and trivalents bound to glass beads stall and rescue microtubule shortening [47]. In the context of an Ndc80 trivalent the CH-domain is required for stable binding to the microtubule lattice, while the presence of the Ndc80 tail and its overall charge contributes to the trivalent’s ability to follow the shortening microtubule ends and to rescue microtubule shortening under force [48].

Centromere protein F

Another example of an efficient coupler is centromere protein F (CENP-F), a component of the corona, a fibrous protein network expanding at the kinetochore at the onset of mitosis [49]. CENP-F regulates interactions between the corona and the microtubules, and is also found at the mitochondria and at the nuclear pores [11,50,51]. Despite its multiple localizations, CENP-F is not essential for viability, and its deletion has a very mild mitotic phenotype [50,52]. CENP-F contains an N-terminal microtubule-binding site with a unique fold and an increased affinity for bent tubulin, and an unstructured positively charged C-terminal tail binding preferentially to straight tubulin [53] (Table 1). When present in multiple copies on the surface of a cargo, both N- and C-terminal MTBDs follow microtubule growth and shortening and capture the force generated by microtubule shortening [10,53].

CENP-E

CENP-E is a potential paralog of CENP-F with a kinesin-like motor domain at the N-terminus and an unstructured positively charged C-terminal tail [49,51] (Table 1). CENP-E is a component of corona and is important for chromosome congression in mitosis [54]. The C-terminal tail is required to retain the CENP-E molecule at the microtubule plus-end if its N-terminal plus-end directed motor domain walks off the microtubule tip [55]. The combination of the motor and the unstructured tail domains allows CENP-E to stay bound to both growing and shortening microtubule ends, but CENP-E’s ability to capture microtubule-generated force has not been reported.

KKT4

Kinetoplastids build a kinetochore that is completely unrelated to any other eukaryote group [56]. However, the principal microtubule-binding protein in trypanosomes, KKT4, shares the pattern with the proteins described above: its microtubule-binding site contains an unstructured positively charged sequence and a folded helical domain (Table 1). This domain combination enables KKT4 to follow shortening microtubule in single-molecule conditions in the absence of force, and against the opposing force when multiple copies of KKT4 are bound to a bead [57].

Potential end-couplers with disordered microtubule-binding sequences

CENP-Q

A component of human kinetochore, was shown to bind microtubules through its N-terminal unstructured sequence [58]. Chromosome congression defects caused by the deletion of CENP-Q1-67 could be rescued by fusion of NDC801-80 instead. These observations suggest functional similarity between the N-terminal tails of Ndc80 and CENP-Q, but it remains to be tested whether CENP-Q can follow microtubule shortening on its own, and whether folded domains contribute to CENP-Q’s interaction with microtubules.

TOG-domain proteins

Conserved globular TOG-domains bind to the very ends of bent tubulin protofilaments at the microtubule tip. TOG-domain proteins enhance microtubule growth, like homologous proteins XMAP215 (Xenopus laevis) and Stu2 (S. cerevisiae) [59,60], or stabilize microtubule ends, like CLASP proteins [61,62]. However, at least in the case of XMAP215, the efficient binding to the microtubule requires an unstructured positively charged linker between TOG4 and TOG5 [63] (Table 1). Beads coated with multiple copies of XMAP215 could be transported by growing microtubule ends against a small opposing force [64], but it is unknown whether XMAP215 captures the force released during microtubule shortening. Stu2, a budding yeast homolog of XMAP215, plays an important role at the kinetochore-microtubule interface by enhancing the sensitivity of purified kinetochores to tension [65].

Another protein in this group is Abl2 tyrosine kinase, which can phosphorylate XMAP215 and CLASP [66,67], and was shown to interact with microtubules directly and sometimes follow the growing microtubule ends [68]. Microtubule-binding of Abl2 requires a central unstructured region that is not positively charged, contrary to all other examples described above. Finally, it should be mentioned that XMAP215 was shown to follow shortening microtubule ends, but it was not shown for CLASP or Abl2, and their force-coupling properties are also unclear.

Microtubule features supporting end-tracking and force capture

The shortening ends of microtubules are characterized by tubulin protofilaments bending outward in a shape resembling ‘ram’s horns’ and generating power strokes as microtubule disassembles [69,70] (Figure 1A). One of the force-coupling models discussed in the literature proposes that the couplers bind bent tubulin with high affinity to transform the power strokes into processive motion of the cargo [21,71] (Figure 3A). Indeed, some of the couplers described above bind bent tubulin preferentially, like Ska, N-terminus of CENP-F or CLASP [20,53]. However, not all proteins that have this property follow microtubule shortening, while many efficient force-couplers bind preferentially to the straight tubulin lattice in bulk biochemical assays, like Ndc80 [20], C-terminus of CENP-F [53] or Dam1, which follows microtubule ends thanks to its increased affinity for GTP-tubulin [72].

Microtubule features supporting end-tracking

Figure 3
Microtubule features supporting end-tracking

(A) A coupler with increased affinity for bent tubulin should be enriched at the ends of the microtubule, where protofilaments are bent. (B) Free diffusion of weakly bound coupler (shown with an arrow) without a specific affinity for microtubule end is biased in the direction of microtubule shortening by the shrinking of the available microtubule lattice. (C) Negative charges of the tubulin C-termini are spatially enriched on the inner side of the bent tubulin protofilaments at the microtubule end. This increased density of negative charges at the microtubule end (shown in cyan) retains positively charged microtubule couplers close to microtubule end (shown in orange).

Figure 3
Microtubule features supporting end-tracking

(A) A coupler with increased affinity for bent tubulin should be enriched at the ends of the microtubule, where protofilaments are bent. (B) Free diffusion of weakly bound coupler (shown with an arrow) without a specific affinity for microtubule end is biased in the direction of microtubule shortening by the shrinking of the available microtubule lattice. (C) Negative charges of the tubulin C-termini are spatially enriched on the inner side of the bent tubulin protofilaments at the microtubule end. This increased density of negative charges at the microtubule end (shown in cyan) retains positively charged microtubule couplers close to microtubule end (shown in orange).

The relation between the affinity for bent tubulin and the end-coupling properties is further complicated by the observations that tubulin protofilaments at the growing microtubule ends are as bent as at the shortening ones [3] (Figure 1). If specific affinity is the only feature necessary for the end tracking, the couplers should follow the growing ends as efficiently as the shortening ones. While this bidirectional tracking has been observed for XMAP215 [59], and to a lesser extent for Ska [23,24], other couplers with preference for bent tubulin are unidirectional: N-terminus of CENP-F moves with the shortening microtubules [53] with rare exceptions [10], while Stu2 and a single TOG-domain of CLASP2 fused to a positively charged linker follow microtubule growth [60,62]. One possible explanation for this discrepancy is given by an almost 10-fold difference in the growth and shortening speeds of microtubules in vitro, which might make the dissociation and re-association rates of the coupling molecules limiting for their processivity.

An alternative model of ‘biased diffusion’ is used to explain how freely diffusing molecules can promote cargo motion with a shortening microtubule end: the microtubule ‘lattice shrinking’ biases net movement of a cargo as the microtubule shortens [21,71] (Figure 3B). This model is attractive because it explains how couplers with no affinity to microtubule ends can still follow microtubule shortening. In the absence of an interaction with the end, the couplers moving by biased diffusion do not influence microtubule dynamics. For multivalent couplers like a full Dam1 ring [28] or an Ndc80 oligomer [47] this model becomes less suitable, because the coupler’s diffusion is negligibly slow compared with the typical lifetime of a dynamic microtubule. Microtubule depolymerization forces the practically immobile coupler either to move or to detach. Consequently, the shortening microtubule end spends a fraction of the released energy to move such a strongly bound coupler, resulting in a slowdown of the shortening speed [34,47,53], contrary to the predictions of the biased diffusion model [38,46].

Many of the couplers discussed in this review rely in their microtubule binding on the unstructured negatively charged C-termini of tubulin, including the disordered C-termini of CENP-F and Dam1, CENP-E tail and Ndc80 [53,73–75], but not globular MTBD of Ska [16]. One of the most studied couplers, Ndc80, is binding preferentially to straight tubulin lattice in bulk [20], but relies on its positively charged N-terminus to stall and rescue microtubule shortening under force as a multivalent [47,48]. Because Ndc80 multivalents stall microtubule shortening after the microtubule end has generated force, i.e. after the power-stroke generated by tubulin protofilaments, this interaction between Ndc80 and microtubule is unlikely to happen along the stable microtubule lattice, away from the action of microtubule-generated force. Ndc80-mediated rescue and stalling of microtubule shortening suggests that Ndc80 multivalents transiently interact with the tubulin protofilaments under force. This hypothesis is further corroborated by an observation that Ndc80-microtubule connection stiffens during stall, with the stiffness and the stalling force positively correlating [47]. A similar effect, although documented with less detail, is observed for MACF2 C-terminus [9]. Theoretical considerations suggest that the stiffness of the microtubule-kinetochore link is an important parameter to rationalize kinetochore oscillations observed in mitotic cells [76]. Further studies will have to address the nature of the tubulin interface at the shortening microtubule end which mediates the force-dependent stiffening connection to multivalent unstructured couplers. It is attractive to speculate that the binding interface could be formed by an increased density of negatively charged tubulin C-termini on the inner side of tubulin ‘ram’s horns’ (Figure 3C). Positively charged sequences could then be enriched in these negatively charged regions long enough (thanks to multivalency and the presence of additional folded MTBDs) to allow microtubule-generated force to be transmitted to the cargo, or for the opposing force from the cargo to stall microtubule shortening and put tubulin protofilaments in a configuration compatible with rescue and regrowth.

Capturing the microtubule-generated force

Shortening microtubules generate force thanks to the energy of GTP hydrolysis stored in the tubulin lattice and released in the form of power strokes of tubulin protofilaments bending outward (Figure 1A). Theoretical predictions estimate the maximum force a microtubule could generate in the range of tens of piconewtons [77–79]. The efficiency of a force coupler can be defined as the amount of microtubule-generated force that is transmitted to the cargo. Another parameter that is used to assess a coupler is load-bearing capacity, or the amount of force the coupler can withstand before its connection to microtubule is lost.

While both force-efficiency and load-bearing capacity are important for the couplers’ performance, they are estimated using different experimental methods with the same readout expressed in pN (Figure 2A), leading to some confusion in the literature. For the purposes of this review, I discuss the experimental methods in their use to estimate the couplers’ force-efficiency. I separate the experimental methods in two groups, one in which a pre-defined force is imposed on the coupler–microtubule connection by the experimenter (Figure 4A–C), and the other in which the force is exerted by microtubule end without any pre-conception of the force magnitude (Figure 4D,E). In all cases however, an integral part of the force-measurement system is a glass or plastic sphere that is held with an optical trap, and interacts with the microtubule through coupling proteins attached to the sphere's surface (Figure 4A,D).

Experimental assays to probe microtubule force-coupling

Figure 4
Experimental assays to probe microtubule force-coupling

(A) Assays with force applied by the experimenter, usually through a piezo-driven microscope stage. Displacement of the coverslip-attached microtubule bound to a bead creates a returning force acting toward the centre of the optical trap. The force can be gradually ramped up at a constant speed until the bead-microtubule connection is ruptured (B). Alternatively, the force can be kept constant through a feedback loop (C, blue line). In a constant force (or ‘force-clamp’) setup the microscope stage is constantly updated to compensate for the bead movement with the microtubule, so the stage movements can serve as a readout for microtubule dynamics under force (C, red line). (D) Assay with force applied by the microtubule end. The optical trap is kept stationary while the bead is displaced by the shortening microtubule end. Bead displacement is calibrated into force. As the microtubule-generated force increases, the bead moves slower against the opposing force from the optical trap, until these forces are equalized and the microtubule shortening is stalled (E). After the stall, the bead can detach and microtubule resumes the shortening, or the bead can remain attached and force the microtubule end to switch to growth (rescue). Both of these alternative outcomes are shown in (E).

Figure 4
Experimental assays to probe microtubule force-coupling

(A) Assays with force applied by the experimenter, usually through a piezo-driven microscope stage. Displacement of the coverslip-attached microtubule bound to a bead creates a returning force acting toward the centre of the optical trap. The force can be gradually ramped up at a constant speed until the bead-microtubule connection is ruptured (B). Alternatively, the force can be kept constant through a feedback loop (C, blue line). In a constant force (or ‘force-clamp’) setup the microscope stage is constantly updated to compensate for the bead movement with the microtubule, so the stage movements can serve as a readout for microtubule dynamics under force (C, red line). (D) Assay with force applied by the microtubule end. The optical trap is kept stationary while the bead is displaced by the shortening microtubule end. Bead displacement is calibrated into force. As the microtubule-generated force increases, the bead moves slower against the opposing force from the optical trap, until these forces are equalized and the microtubule shortening is stalled (E). After the stall, the bead can detach and microtubule resumes the shortening, or the bead can remain attached and force the microtubule end to switch to growth (rescue). Both of these alternative outcomes are shown in (E).

Assays with pre-defined force

In a ‘rupture force’ assay the optically trapped and microtubule-bound bead is subjected to a linearly increasing force until the bead-microtubule attachment breaks (Figure 4A,B). This assay has been used in a number of publications to probe the load-bearing of reconstituted kinetochore complexes [18,80–82] or whole kinetochore particles purified from cells [65,83]. This assay is attractive for its simplicity, however, as long as the amount of exerted force is decided by the experimenter, the rupture force assay is not suitable to probe the couplers’ ability to capture microtubule-generated force. In a variation of this approach, the force acting on the bead is kept constant while the bead is following the growing and shortening microtubule ends. Constant force, or ‘force-clamp’ was very valuable to assess the effect of tension on microtubule dynamic parameters, such as the shortening speed or the frequency of switches from growth to shortening (a catastrophe), and from shortening to growth (a rescue) [18,65,83] (Figure 4C).

Assays with microtubule-generated force

To address the coupler’s force-transmission experimentally, a shortening microtubule is made to pull on an optically trapped bead coated with coupling molecules; displacement of the bead is calibrated to estimate the force (Figure 4D,E). This assay is characterized by a stationary trap that does not follow microtubule dynamics, like in force-clamp assay. A predictor of the correct estimation of microtubule-generated force is stalling, an equilibrium situation in which the microtubule-generated force is at balance with the returning force exerted by the optical trap on the bead (Figure 4F). The stall can result either in a detachment of the bead from the microtubule, or if there is soluble tubulin present in the assay, in force-dependent rescue [47,48] (Figure 4E). This approach allowed to identify the unstructured N-terminal tail of Ndc80 as the main contributor to the duration of the microtubule stalling by Ndc80 oligomers, and the duration of the stall as a predictor of rescue [48]. Force-clamp assay is also suitable to estimate microtubule stall force by finding the minimal applied force that prevents movement of otherwise processive cargo [83]. However, in the stationary trap assay (Figure 4D,E) the whole range of microtubule-generated forces and coupler movement speeds can be sampled from one measurement [39].

Concluding remarks

Given the abundance of disordered domains in the proteome (reviewed in [84]), specific requirements for a disordered domain to become a microtubule coupler remain to be found. This question is especially interesting because of very poor sequence conservation among the disordered microtubule couplers. Even the N-terminal tail of the evolutionary conserved Ndc80 complex is only poorly conserved across its orthologs [85]. In light of rapid evolution of disordered proteins compared with folded ones (reviewed in [86]) there are open interesting questions: does a specific amino acid sequence matter to make an efficient microtubule coupler? Do coupler sequences co-evolve with tubulin C-terminal tails? Do specific coupler sequences co-occur with specific tubulin post-translational modifications in different cell types or cell cycle phases?

Summary

  • Majority of proteins following shortening microtubule ends act as multimers and rely on unstructured positively charged sequences in this activity.

  • The unstructured microtubule-biding domains are not conserved in their sequence, but share overall positive charge and amino acid composition.

  • High affinity for bent tubulin is not necessary for end-tracking and capture of microtubule-generated force.

  • Microtubule stalling and rescue happens through a poorly understood interaction that might stiffen under force.

Competing Interests

The author declares that there are no competing interests associated with the manuscript.

Funding

This work was supported by the European Research Council Synergy Grant MODELCELL [grant number 609822 (to Marileen Dogterom and Anna Akhmanova)].

Acknowledgements

I am grateful to Nikita Gudimchuk for sharing the microtubule coordinates, and to Nemo Andrea for the 3D rendering of these coordinates to produce Figure 1A. I am also grateful to Nikita Gudimchuk, Pim Huis in șt Veld, Natalia Kochergina and Anna Akhmanova for comments on the manuscript and valuable discussions.

Abbreviations

     
  • Catastrophe

    a stochastic switch from microtubule growth to shortening

  •  
  • CENP

    centromere protein

  •  
  • CH

    calponin homology

  •  
  • Coupler

    a protein complex/oligomer that passively binds to microtubule and utilizes microtubule-generated force to move itself or a cargo

  •  
  • End-tracking

    ability of a coupler to retain its association to the end of the microtubule as the microtubule shortens. The length/duration of end-tracking is also referred to as processivity

  •  
  • ER

    endoplasmic reticulum

  •  
  • MACF2

    microtubule-actin cross-linking factor 2

  •  
  • MTBD

    microtubule-binding domain

  •  
  • Multivalency

    coupler’s ability to bind microtubules through multiple (similar or different) MTBDs

  •  
  • Optical trap

    an instrument to measure forces in the piconewton (pN) range. Features a parallel beam of an infrared laser focused by a microscope objective into a diffraction-limited spot. Glass or plastic spheres (beads) are attracted to the centre of the focused beam, and a force is necessary to displace the bead from the trap centre

  •  
  • Protofilament

    linear strand of αβ-tubulin dimers attached head to tail

  •  
  • Rescue

    a stochastic switch from microtubule shortening to growth

  •  
  • Stall

    microtubule shortening that is forcefully slowed down to a pause

References

References
1.
McIntosh
J.R.
,
Volkov
V.
,
Ataullakhanov
F.I.
and
Grishchuk
E.L.
(
2010
)
Tubulin depolymerization may be an ancient biological motor
.
J. Cell. Sci.
123
,
3425
3434
[PubMed]
2.
Brouhard
G.J.
and
Rice
L.M.
(
2018
)
Microtubule dynamics: an interplay of biochemistry and mechanics
.
Nat. Rev. Mol. Cell. Biol.
19
,
451
463
[PubMed]
3.
McIntosh
J.R.
,
O’Toole
E.
,
Morgan
G.
,
Austin
J.
,
Ulyanov
E.
,
Ataullakhanov
F.
et al.
(
2018
)
Microtubules grow by the addition of bent guanosine triphosphate tubulin to the tips of curved protofilaments
.
J. Cell Biol.
217
,
2691
2708
[PubMed]
4.
Zhang
R.
,
LaFrance
B.
and
Nogales
E.
(
2018
)
Separating the effects of nucleotide and EB binding on microtubule structure
.
Proc. Natl. Acad. Sci. U.S.A.
115
,
E6191
E6200
[PubMed]
5.
Inoue
S.
and
Sato
H.
(
1967
)
Cell motility by labile association of molecules. The nature of mitotic spindle fibers and their role in chromosome movement
.
J. Gen. Physiol.
50
,
259
292
6.
Vleugel
M.
,
Kok
M.
and
Dogterom
M.
(
2016
)
Understanding force-generating microtubule systems through in vitro reconstitution
.
Cell. Adh. Migr.
10
,
475
494
[PubMed]
7.
Waterman-Storer
C.M.
,
Gregory
J.
,
Parsons
S.F.
and
Salmon
E.D.
(
1995
)
Membrane/microtubule tip attachment complexes (TACs) allow the assembly dynamics of plus ends to push and pull membranes into tubulovesicular networks in interphase Xenopus egg extracts
.
J. Cell Biol.
130
,
1161
1169
[PubMed]
8.
Guo
Y.
,
Li
D.
,
Zhang
S.
,
Yang
Y.
,
Liu
J.J.
,
Wang
X.
et al.
(
2018
)
Visualizing intracellular organelle and cytoskeletal interactions at nanoscale resolution on millisecond timescales
.
Cell
175
,
1430
1442.e17
9.
Rodríguez-García
R.
,
Volkov
V.A.
,
Chen
C.-Y.
,
Katrukha
E.A.
,
Olieric
N.
,
Aher
A.
et al.
(
2020
)
Mechanisms of motor-independent membrane remodeling driven by dynamic microtubules
.
Curr. Biol.
30
,
972
987.e12
10.
Kanfer
G.
,
Peterka
M.
,
Arzhanik
V.K.
,
Drobyshev
A.L.
,
Ataullakhanov
F.I.
,
Volkov
V.A.
et al.
(
2017
)
CENP-F couples cargo to growing and shortening microtubule ends
.
Mol. Biol. Cell
28
,
2400
2409
[PubMed]
11.
Kanfer
G.
,
Courtheoux
T.
,
Peterka
M.
,
Meier
S.
,
Soste
M.
,
Melnik
A.
et al.
(
2015
)
Mitotic redistribution of the mitochondrial network by Miro and Cenp-F
.
Nat. Commun.
6
,
8015
[PubMed]
12.
Koshland
D.E.
,
Mitchison
T.J.
and
Kirschner
M.W.
(
1988
)
Polewards chromosome movement driven by microtubule depolymerization in vitro
.
Nature
331
,
499
504
[PubMed]
13.
Jones
D.T.
and
Cozzetto
D.
(
2015
)
DISOPRED3: precise disordered region predictions with annotated protein-binding activity
.
Bioinformatics
31
,
857
863
[PubMed]
14.
Hanisch
A.
,
Silljé
H.H.
and
Nigg
E.A.
(
2006
)
Timely anaphase onset requires a novel spindle and kinetochore complex comprising Ska1 and Ska2
.
EMBO J.
25
,
5504
5515
[PubMed]
15.
Raaijmakers
J.A.
,
Tanenbaum
M.E.
,
Maia
A.F.
and
Medema
R.H.
(
2009
)
RAMA1 is a novel kinetochore protein involved in kinetochore-microtubule attachment
.
J. Cell Sci.
122
,
2436
2445
[PubMed]
16.
Abad
M.A.
,
Medina
B.
,
Santamaria
A.
,
Zou
J.
,
Plasberg-Hill
C.
,
Madhumalar
A.
et al.
(
2014
)
Structural basis for microtubule recognition by the human kinetochore Ska complex
.
Nat. Commun.
5
,
2964
[PubMed]
17.
Redli
P.M.
,
Gasic
I.
,
Meraldi
P.
,
Nigg
E.A.
and
Santamaria
A.
(
2016
)
The Ska complex promotes Aurora B activity to ensure chromosome biorientation
.
J. Cell Biol.
215
,
77
93
[PubMed]
18.
Helgeson
L.A.
,
Zelter
A.
,
Riffle
M.
,
MacCoss
M.J.
,
Asbury
C.L.
and
Davis
T.N.
(
2018
)
Human Ska complex and Ndc80 complex interact to form a load-bearing assembly that strengthens kinetochore-microtubule attachments
.
Proc. Natl. Acad. Sci. U.S.A.
115
,
2740
2745
[PubMed]
19.
Abad
M.A.
,
Zou
J.
,
Medina-Pritchard
B.
,
Nigg
E.A.
,
Rappsilber
J.
,
Santamaria
A.
et al.
(
2016
)
Ska3 ensures timely mitotic progression by interacting directly with microtubules and Ska1 microtubule binding domain
.
Sci. Rep.
6
,
34042
[PubMed]
20.
Schmidt
J.C.
,
Arthanari
H.
,
Boeszoermenyi
A.
,
Dashkevich
N.M.
,
Wilson-Kubalek
E.M.
,
Monnier
N.
et al.
(
2012
)
The kinetochore-bound Ska1 complex tracks depolymerizing microtubules and binds to curved protofilaments
.
Dev. Cell
23
,
968
980
[PubMed]
21.
Grishchuk
E.L.
(
2017
)
Biophysics of Microtubule End Coupling at the Kinetochore
. In
Centromeres and Kinetochores: Discovering the Molecular Mechanisms Underlying Chromosome Inheritance
(
Black
B.E.
, ed.), pp.
397
428
,
Springer International Publishing
,
Cham
22.
Welburn
J.P.I.
,
Grishchuk
E.L.
,
Backer
C.B.
,
Wilson-Kubalek
E.M.
,
Yates
J.R.
and
Cheeseman
I.M.
(
2009
)
The human kinetochore Ska1 complex facilitates microtubule depolymerization-coupled motility
.
Dev. Cell.
16
,
374
385
[PubMed]
23.
Monda
J.K.
,
Whitney
I.P.
,
Tarasovetc
E.V.
,
Wilson-Kubalek
E.
,
Milligan
R.A.
,
Grishchuk
E.L.
et al.
(
2017
)
Microtubule tip tracking by the spindle and kinetochore protein Ska1 requires diverse tubulin-interacting surfaces
.
Curr. Biol.
27
,
3666
3675.e6
24.
Maciejowski
J.
,
Drechsler
H.
,
Grundner-Culemann
K.
,
Ballister
E.R.
,
Rodriguez-Rodriguez
J.A.
,
Rodriguez-Bravo
V.
et al.
(
2017
)
Mps1 regulates kinetochore-microtubule attachment stability via the Ska complex to ensure error-free chromosome segregation
.
Dev. Cell
41
,
143
156.e6
25.
Westermann
S.
,
Wang
H.-W.
,
Avila-Sakar
A.
,
Drubin
D.G.
,
Nogales
E.
and
Barnes
G.
(
2006
)
The Dam1 kinetochore ring complex moves processively on depolymerizing microtubule ends
.
Nature
440
,
565
569
[PubMed]
26.
Cheeseman
I.M.
,
Brew
C.
,
Wolyniak
M.
,
Desai
A.
,
Anderson
S.
,
Muster
N.
et al.
(
2001
)
Implication of a novel multiprotein Dam1p complex in outer kinetochore function
.
J. Cell Biol.
155
,
1137
1146
[PubMed]
27.
Miranda
J.J.L.
,
De Wulf
P.
,
Sorger
P.K.
and
Harrison
S.C.
(
2005
)
The yeast DASH complex forms closed rings on microtubules
.
Nat. Struct. Mol. Biol.
12
,
138
143
28.
Grishchuk
E.L.
,
Spiridonov
I.S.
,
Volkov
V.A.
,
Efremov
A.
,
Westermann
S.
,
Drubin
D.
et al.
(
2008
)
Different assemblies of the DAM1 complex follow shortening microtubules by distinct mechanisms
.
Proc. Natl. Acad. Sci. U.S.A.
105
,
6918
623
[PubMed]
29.
van Hooff
J.J.E.
,
Snel
B.
and
Kops
G.J.P.L.
(
2017
)
Unique phylogenetic distributions of the Ska and Dam1 complexes support functional analogy and suggest multiple parallel displacements of Ska by Dam1
.
Genome Biol. Evol.
9
,
1295
1303
[PubMed]
30.
Wang
H.-W.
,
Ramey
V.H.
,
Westermann
S.
,
Leschziner
A.E.
,
Welburn
J.P.I.
,
Nakajima
Y.
et al.
(
2007
)
Architecture of the Dam1 kinetochore ring complex and implications for microtubule-driven assembly and force-coupling mechanisms
.
Nat. Struct. Mol. Biol.
14
,
721
726
31.
Miranda
J.J.L.
,
King
D.S.
and
Harrison
S.C.
(
2007
)
Protein arms in the kinetochore-microtubule interface of the yeast DASH complex
.
Mol. Biol. Cell
18
,
2503
2510
[PubMed]
32.
Legal
T.
,
Zou
J.
,
Sochaj
A.
,
Rappsilber
J.
and
Welburn
J.P.I.
(
2016
)
Molecular architecture of the Dam1 complex-microtubule interaction
.
Open Biol.
6
,
[PubMed]
33.
Ramey
V.H.
,
Wong
A.
,
Fang
J.
,
Howes
S.
,
Barnes
G.
and
Nogales
E.
(
2011
)
Subunit organization in the Dam1 kinetochore complex and its ring around microtubules
.
Mol. Biol. Cell
22
,
4335
4342
[PubMed]
34.
Grishchuk
E.L.
,
Efremov
A.K.
,
Volkov
V.A.
,
Spiridonov
I.S.
,
Gudimchuk
N.
,
Westermann
S.
et al.
(
2008
)
The Dam1 ring binds microtubules strongly enough to be a processive as well as energy-efficient coupler for chromosome motion
.
Proc. Natl. Acad. Sci. U.S.A.
105
,
15423
15428
[PubMed]
35.
Jenni
S.
and
Harrison
S.C.
(
2018
)
Structure of the DASH/Dam1 complex shows its role at the yeast kinetochore-microtubule interface
.
Science
360
,
552
[PubMed]
36.
Leung
C.L.
,
Sun
D.
,
Zheng
M.
,
Knowles
D.R.
and
Liem
R.K.H.
(
1999
)
Microtubule actin cross-linking factor (Macf): a hybrid of dystonin and dystrophin that can interact with the actin and microtubule cytoskeletons
.
J. Cell Biol.
147
,
1275
1286
[PubMed]
37.
Drechsler
H.
,
Xu
Y.
,
Geyer
V.F.
,
Zhang
Y.
and
Diez
S.
(
2019
)
Multivalent electrostatic microtubule interactions of synthetic peptides are sufficient to mimic advanced MAP-like behavior
.
Mol. Biol. Cell
30
,
2953
2968
[PubMed]
38.
Efremov
A.
,
Grishchuk
E.L.
,
McIntosh
J.R.
and
Ataullakhanov
F.I.
(
2007
)
In search of an optimal ring to couple microtubule depolymerization to processive chromosome motions
.
Proc. Natl. Acad. Sci. U.S.A.
104
,
19017
19022
[PubMed]
39.
Volkov
V.A.
,
Zaytsev
A.V.
,
Gudimchuk
N.
,
Grissom
P.M.
,
Gintsburg
A.L.
,
Ataullakhanov
F.I.
et al.
(
2013
)
Long tethers provide high-force coupling of the Dam1 ring to shortening microtubules
.
Proc. Natl. Acad. Sci. U.S.A.
110
,
7708
7713
[PubMed]
40.
Cheeseman
I.M.
,
Chappie
J.S.
,
Wilson-Kubalek
E.M.
and
Desai
A.
(
2006
)
The conserved KMN network constitutes the core microtubule-binding site of the kinetochore
.
Cell
127
,
983
997
[PubMed]
41.
DeLuca
J.G.
,
Gall
W.E.
,
Ciferri
C.
,
Cimini
D.
,
Musacchio
A.
and
Salmon
E.D.
(
2006
)
Kinetochore microtubule dynamics and attachment stability are regulated by Hec1
.
Cell
127
,
969
982
[PubMed]
42.
Guimaraes
G.J.
,
Dong
Y.
,
McEwen
B.F.
and
DeLuca
J.G.
(
2008
)
Kinetochore-microtubule attachment relies on the disordered N-terminal tail domain of Hec1
.
Curr. Biol.
18
,
1778
1784
[PubMed]
43.
Miller
S.A.
,
Johnson
M.L.
and
Stukenberg
P.T.
(
2008
)
Kinetochore attachments require an interaction between unstructured tails on microtubules and Ndc80(Hec1)
.
Curr. Biol.
18
,
1785
1791
[PubMed]
44.
Zaytsev
A.V.
,
Mick
J.E.
,
Maslennikov
E.
,
Nikashin
B.
,
DeLuca
J.G.
and
Grishchuk
E.L.
(
2015
)
Multisite phosphorylation of the NDC80 complex gradually tunes its microtubule-binding affinity
.
Mol. Biol. Cell
26
,
1829
1844
[PubMed]
45.
McIntosh
J.R.
,
Grishchuk
E.L.
,
Morphew
M.K.
,
Efremov
A.K.
,
Zhudenkov
K.
,
Volkov
V.A.
et al.
(
2008
)
Fibrils connect microtubule tips with kinetochores: a mechanism to couple tubulin dynamics to chromosome motion
.
Cell
135
,
322
333
[PubMed]
46.
Powers
A.F.
,
Franck
A.D.
,
Gestaut
D.R.
,
Cooper
J.
,
Gracyzk
B.
,
Wei
R.R.
et al.
(
2009
)
The Ndc80 kinetochore complex forms load-bearing attachments to dynamic microtubule tips via biased diffusion
.
Cell
136
,
865
875
[PubMed]
47.
Volkov
V.A.
,
Huis in ’t Veld
P.J.
,
Dogterom
M.
and
Musacchio
A.
(
2018
)
Multivalency of NDC80 in the outer kinetochore is essential to track shortening microtubules and generate forces
.
Elife
7
,
[PubMed]
48.
Huis in ’t Veld
P.J.
,
Volkov
V.A.
,
Stender
I.D.
,
Musacchio
A.
and
Dogterom
M.
(
2019
)
Molecular determinants of the Ska-Ndc80 interaction and their influence on microtubule tracking and force-coupling
.
Elife
8
,
[PubMed]
49.
Ciossani
G.
,
Overlack
K.
,
Petrovic
A.
,
Huis in ’t Veld
P.J.
,
Koerner
C.
,
Wohlgemuth
S.
et al.
(
2018
)
The kinetochore proteins CENP-E and CENP-F directly and specifically interact with distinct BUB mitotic checkpoint Ser/Thr kinases
.
J. Biol. Chem.
293
,
10084
10101
[PubMed]
50.
Auckland
P.
,
Roscioli
E.
,
Coker
H.L.E.
and
McAinsh
A.D.
(
2020
)
CENP-F stabilizes kinetochore-microtubule attachments and limits dynein stripping of corona cargoes
.
J. Cell Biol.
219
,
[PubMed]
51.
Berto
A.
,
Yu
J.
,
Morchoisne-Bolhy
S.
,
Bertipaglia
C.
,
Vallee
R.
,
Dumont
J.
et al.
(
2018
)
Disentangling the molecular determinants for Cenp-F localization to nuclear pores and kinetochores
.
EMBO Rep.
19
,
e44742
[PubMed]
52.
Peterka
M.
and
Kornmann
B.
(
2019
)
Miro-dependent mitochondrial pool of CENP-F and its farnesylated C-terminal domain are dispensable for normal development in mice
.
PLoS Genet.
15
,
e1008050
[PubMed]
53.
Volkov
V.A.
,
Grissom
P.M.
,
Arzhanik
V.K.
,
Zaytsev
A.V.
,
Renganathan
K.
,
McClure-Begley
T.
et al.
(
2015
)
Centromere protein F includes two sites that couple efficiently to depolymerizing microtubules
.
J. Cell Biol.
209
,
813
828
[PubMed]
54.
Kapoor
T.M.
,
Lampson
M.A.
,
Hergert
P.
,
Cameron
L.
,
Cimini
D.
,
Salmon
E.D.
et al.
(
2006
)
Chromosomes can Congress to the metaphase plate before biorientation
.
Science
311
,
388
[PubMed]
55.
Gudimchuk
N.
,
Vitre
B.
,
Kim
Y.
,
Kiyatkin
A.
,
Cleveland
D.W.
,
Ataullakhanov
F.I.
et al.
(
2013
)
Kinetochore kinesin CENP-E is a processive bi-directional tracker of dynamic microtubule tips
.
Nat. Cell Biol.
15
,
1079
1088
[PubMed]
56.
Akiyoshi
B.
and
Gull
K.
(
2014
)
Discovery of unconventional kinetochores in kinetoplastids
.
Cell
156
,
1247
1258
[PubMed]
57.
Llauro
A.
,
Hayashi
H.
,
Bailey
M.E.
,
Wilson
A.
,
Ludzia
P.
,
Asbury
C.L.
et al.
(
2018
)
The kinetoplastid kinetochore protein KKT4 is an unconventional microtubule tip-coupling protein
.
J. Cell Biol.
217
,
3886
3900
[PubMed]
58.
Pesenti
M.E.
,
Prumbaum
D.
,
Auckland
P.
,
Smith
C.M.
,
Faesen
A.C.
,
Petrovic
A.
et al.
(
2018
)
Reconstitution of a 26-subunit human kinetochore reveals cooperative microtubule binding by CENP-OPQUR and NDC80
.
Mol. Cell
71
,
923
939.e10
59.
Brouhard
G.J.
,
Stear
J.H.
,
Noetzel
T.L.
,
Al-Bassam
J.
,
Kinoshita
K.
,
Harrison
S.C.
et al.
(
2008
)
XMAP215 is a processive microtubule polymerase
.
Cell
132
,
79
88
[PubMed]
60.
Podolski
M.
,
Mahamdeh
M.
and
Howard
J.
(
2014
)
Stu2, the budding yeast XMAP215/Dis1 homolog, promotes assembly of yeast microtubules by increasing growth rate and decreasing catastrophe frequency
.
J. Biol. Chem.
289
,
28087
28093
[PubMed]
61.
Al-Bassam
J.
,
Kim
H.
,
Brouhard
G.
,
van Oijen
A.
,
Harrison
S.C.
and
Chang
F.
(
2010
)
CLASP promotes microtubule rescue by recruiting tubulin dimers to the microtubule
.
Dev. Cell
19
,
245
258
[PubMed]
62.
Aher
A.
,
Kok
M.
,
Sharma
A.
,
Rai
A.
,
Olieric
N.
,
Rodriguez-Garcia
R.
et al.
(
2018
)
CLASP suppresses microtubule catastrophes through a single TOG domain
.
Dev. Cell
46
,
40
58.e8
63.
Widlund
P.O.
,
Stear
J.H.
,
Pozniakovsky
A.
,
Zanic
M.
,
Reber
S.
,
Brouhard
G.J.
et al.
(
2011
)
XMAP215 polymerase activity is built by combining multiple tubulin-binding TOG domains and a basic lattice-binding region
.
Proc. Natl. Acad. Sci. U.S.A.
108
,
2741
2746
[PubMed]
64.
Trushko
A.
,
Schäffer
E.
and
Howard
J.
(
2013
)
The growth speed of microtubules with XMAP215-coated beads coupled to their ends is increased by tensile force
.
Proc. Natl. Acad. Sci. U.S.A.
110
,
14670
65.
Miller
M.P.
,
Asbury
C.L.
and
Biggins
S.
(
2016
)
A TOG protein confers tension sensitivity to kinetochore-microtubule attachments
.
Cell
165
,
1428
1439
[PubMed]
66.
Engel
U.
,
Zhan
Y.
,
Long
J.B.
,
Boyle
S.N.
,
Ballif
B.A.
,
Dorey
K.
et al.
(
2014
)
Abelson phosphorylation of CLASP2 modulates its association with microtubules and actin
.
Cytoskeleton
71
,
195
209
[PubMed]
67.
Martin
M.
,
Ahern-Djamali
S.M.
,
Hoffmann
F.M.
and
Saxton
W.M.
(
2005
)
Abl tyrosine kinase and its substrate Ena/VASP have functional interactions with kinesin-1
.
Mol. Biol. Cell.
16
,
4225
4230
[PubMed]
68.
Hu
Y.
,
Lyu
W.
,
Lowery
L.A.
and
Koleske
A.J.
(
2019
)
Regulation of MT dynamics via direct binding of an Abl family kinase
.
J. Cell Biol.
218
,
3986
3997
[PubMed]
69.
Grishchuk
E.L.
,
Molodtsov
M.I.
,
Ataullakhanov
F.I.
and
McIntosh
J.R.
(
2005
)
Force production by disassembling microtubules
.
Nature
438
,
384
388
[PubMed]
70.
Driver
J.W.
,
Geyer
E.A.
,
Bailey
M.E.
,
Rice
L.M.
and
Asbury
C.L.
(
2017
)
Direct measurement of the force-generating capacity of protofilaments curling outward from disassembling microtubule tips
.
Biophys. J.
112
,
565a
565a
71.
Asbury
C.L.
,
Tien
J.F.
and
Davis
T.N.
(
2011
)
Kinetochores’ gripping feat: conformational wave or biased diffusion?
Trends Cell Biol.
21
,
38
46
[PubMed]
72.
Lampert
F.
,
Hornung
P.
and
Westermann
S.
(
2010
)
The Dam1 complex confers microtubule plus end-tracking activity to the Ndc80 kinetochore complex
.
J. Cell Biol.
189
,
641
649
[PubMed]
73.
Ramey
V.H.
,
Wang
H.W.
,
Nakajima
Y.
,
Wong
A.
,
Liu
J.
,
Drubin
D.
et al.
(
2011
)
The Dam1 ring binds to the E-hook of tubulin and diffuses along the microtubule
.
Mol. Biol. Cell.
22
,
457
466
[PubMed]
74.
Musinipally
V.
,
Howes
S.
,
Alushin
G.M.
and
Nogales
E.
(
2013
)
The microtubule binding properties of CENP-E's C-terminus and CENP-F
.
J. Mol. Biol.
425
,
4427
4441
[PubMed]
75.
Ciferri
C.
,
Pasqualato
S.
,
Screpanti
E.
,
Varetti
G.
,
Santaguida
S.
,
Dos Reis
G.
et al.
(
2008
)
Implications for kinetochore-microtubule attachment from the structure of an engineered Ndc80 complex
.
Cell
133
,
427
439
[PubMed]
76.
Schwietert
F.
and
Kierfeld
J.
(
2020
)
Bistability and oscillations in cooperative microtubule and kinetochore dynamics in the mitotic spindle
.
New J. Phys.
22
,
053008
77.
Vichare
S.
,
Jain
I.
,
Inamdar
M.M.
and
Padinhateeri
R.
(
2013
)
Forces due to curving protofilaments in microtubules
.
Phys. Rev. E
88
,
062708
78.
Molodtsov
M.I.
,
Grishchuk
E.L.
,
Efremov
A.K.
,
McIntosh
J.R.
and
Ataullakhanov
F.I.
(
2005
)
Force production by depolymerizing microtubules: a theoretical study
.
Proc. Natl. Acad. Sci. U.S.A.
102
,
4353
4358
[PubMed]
79.
Joglekar
A.P.
and
Hunt
A.J.
(
2002
)
A simple, mechanistic model for directional instability during mitotic chromosome movements
.
Biophys. J.
83
,
42
58
[PubMed]
80.
Umbreit
N.T.
,
Miller
M.P.
,
Tien
J.F.
,
Ortola
J.C.
,
Gui
L.
,
Lee
K.K.
et al.
(
2014
)
Kinetochores require oligomerization of Dam1 complex to maintain microtubule attachments against tension and promote biorientation
.
Nat. Commun.
5
,
4951
[PubMed]
81.
Tien
J.F.
,
Umbreit
N.T.
,
Zelter
A.
,
Riffle
M.
,
Hoopmann
M.R.
,
Johnson
R.S.
et al.
(
2014
)
Kinetochore biorientation in Saccharomyces cerevisiae requires a tightly folded conformation of the Ndc80 complex
.
Genetics
198
,
1483
1493
[PubMed]
82.
Tien
J.F.
,
Umbreit
N.T.
,
Gestaut
D.R.
,
Franck
A.D.
,
Cooper
J.
,
Wordeman
L.
et al.
(
2010
)
Cooperation of the Dam1 and Ndc80 kinetochore complexes enhances microtubule coupling and is regulated by Aurora B
.
J. Cell Biol.
189
,
713
723
[PubMed]
83.
Akiyoshi
B.
,
Sarangapani
K.K.
,
Powers
A.F.
,
Nelson
C.R.
,
Reichow
S.L.
,
Arellano-Santoyo
H.
et al.
(
2010
)
Tension directly stabilizes reconstituted kinetochore-microtubule attachments
.
Nature
468
,
576
579
[PubMed]
84.
Dunker
A.K.
,
Silman
I.
,
Uversky
V.N.
and
Sussman
J.L.
(
2008
)
Function and structure of inherently disordered proteins
.
Curr. Opin. Struct. Biol.
18
,
756
764
[PubMed]
85.
Alushin
G.M.
,
Musinipally
V.
,
Matson
D.
,
Tooley
J.
,
Stukenberg
P.T.
and
Nogales
E.
(
2012
)
Multimodal microtubule binding by the Ndc80 kinetochore complex
.
Nat. Struct. Mol. Biol.
19
,
1161
1167
[PubMed]
86.
Brown
C.J.
,
Johnson
A.K.
,
Dunker
A.K.
and
Daughdrill
G.W.
(
2011
)
Evolution and disorder
.
Curr. Opin. Struct. Biol.
21
,
441
446
[PubMed]